AIX-MARSEILLE UNIVERSITE ECOLE DOCTORALE DES SCIENCES DE LA VIE ET DE LA SANTE FACULTE DE MEDECINE DE MARSEILLE Unité de Recherche Microbes, Evolution, Phylogeny and Infection (MEФI)
IHU-Méditerranée infection
Thèse présentée pour obtenir le grade de Doctorat d’Aix-Marseille Université
Spécialité Pathologie Humaine : Maladies infectieuses
Mme Nadia AMANZOUGAGHENE -MEHALLA
Resistance et évolution des poux humains Pediculus humanus
Soutenue le 05 Juillet 2018 devant le jury :
Mme le Professeur Fabienne BREGEON Présidente du jury Mr le Docteur Arezki IZRI Rapporteur Mr le Professeur Lionel ZENNER Rapporteur Mr le Docteur Oleg MEDIANNIKOV Co-directeur de thèse
Direction de la Thèse :
Mr le Professeur Didier RAOULT Directeur de thèse Mr le Docteur Oleg MEDIANNIKOV Co-directeur de thèse
Année Universitaire 2017-2018
Remerciements
Je commencerai par remercier le Professeur Didier RAOULT pour m’avoir accueilli au sein de son laboratoire et permis de réaliser ces travaux de recherche sous sa direction et ses conseils avisés. Permettez-moi de vous exprimer mon profond respect.
Mes profonds remerciements au Docteur Oleg MEDIANNIKOV pour avoir dirigé ce travail.
Vous m’avez donné un support scientifique incontournable et la liberté d’évoluer comme un chercheur. Merci pour m’avoir donné votre temps, votre patience, votre assistance, vos conseils et surtout votre confiance dans toutes les étapes de ce travail. Aussi grande que puisse être ma gratitude, soyez assuré qu'elle ne sera jamais à la hauteur de tous les efforts que vous avez déployé, je vous témoigne le plus profond de mes plaisirs de travailler avec vous.
Mes sincères et chaleureux remerciements vont au Professeur Florence FENOLLAR, pour avoir co-dirigé ce travail. Votre aide scientifique inestimable, votre soutien moral, votre compréhension et votre gentillesse m’ont beaucoup marqué. J’espère que ce modeste travail témoigne de ma profonde reconnaissance et de ma haute considération.
Je tiens à remercier très chaleureusement le Docteur vétérinaire Bernard DAVOUST pour son soutien et son aide précieuse. Trouvez ici, l'expression de ma profonde gratitude.
Je remercie sincèrement Madame le Professeur Fabienne
BREGEON d’avoir accepté d’examiner et de présider mon jury de thèse.
Je remercie très sincèrement le Docteur Arezki IZRI et le Professeur Lionel ZENNER d’avoir accepté d’évaluer ce travail. Veuillez trouver ici l’expression de ma profonde reconnaissance.
J’adresse mes vifs remerciements et ma profonde reconnaissance au Docteur Idir BITAM pour son aide et pour la confiance dont il m’a investi en me proposant de rejoindre l’IHU- méditerranée infection (ex-URMITE).
Je remercie tous mes co-auteurs que je ne pourrais citer individuellement car ils sont nombreux.
Mes sincères remerciements s’adressent à Jean-Michel BERENGER, Yassina BECHAH,
Amira BEN AMARA, Claude NAPPEZ, Pascal WEBER et El Hadji Amadou NIANG, pour leurs conseils et leurs aides précieuses.
Mes plus sincères remerciements vont à l’ensemble des étudiants, technicien(ne)s et au personnel administratif de laboratoire. Plus particulièrement Annick ABEILLE, Elsa
PRUDENT, Emilie, Anne-Marie, François et mes très chères Francine VERIN et Micheline
PITACCOLO pour leurs disponibilités, leurs conseils pratiques et surtout leur soutien moral et leur sourire permanent.
Dédicace
À mes très chers parents, ma mère et mon père, qui par leurs encouragements et leur disponibilité, m’ont poussé à persévérer et à donner le mieux de moi-même Qu’ils trouvent ici le témoignage de ma profonde gratitude.
À mon très cher mari, Messaoud. Merci pour ton soutien indéfectible. Tu as toujours suivi mes études avec intérêt et tu n’as ménagé aucun effort pour ma réussite. Que l’Eternel te garde dans sa bonté.
À mes frères, à mes sœurs, à mes belles-sœurs et beaux-frères pour leur soutien et bienveillance Qu'ils trouvent ici les sentiments de ma reconnaissance.
À mes très chers neveux et nièces Que ce travail vous serve d'exemple pour vous surpasser.
À mes beaux-parents, belles-sœurs et beau-frère pour leur gentillesse Qu'ils trouvent ici les sentiments de ma reconnaissance.
À toute ma famille.
Sommaire
Avant-propos ...... 1
Résumé ...... 2
Abstract ...... 4
Introduction générale ...... 6
Article 1. Revue : Where are we with human lice? A review of the current state of our knowledge...... 10
Chapitre I : Phylogénie et phylogéographie des poux anciens et contemporains ...... 44
Article 2 : High Ancient Genetic Diversity of Human Lice, Pediculus humanus, from Israel Reveals New Insights into the Origin of Clade B Lice...... 46 Article 3 : Mitochondrial diversity and phylogeography of Pediculus humanus, with the description of a new Amazonian Clade F ...... 61
Chapitre II : Epidémiologie des poux et pathogènes associés ...... 91
Article 4 : Head Lice of Pygmies Reveal the Presence of Relapsing Fever Borreliae in the Republic of Congo...... 93 Article 5 : Detection of several emerging bacterial pathogens in human head lice from Mali ...... 113 Article 6 : Molecular survey of Head and Body lice, Pediculus humanus, in France ...... 134 Article 7 : Detection of bacterial pathogens in clade E head lice collected from Niger's refugees in Algeria...... 144 Article 8 : Molecular survey of head lice, Pediculus humanus capitis, in Democratic Republic of Congo ...... 156
Chapitre III : Mécanismes de résistance des poux à l’ivermectine ...... 180 Article 9 : Mutations in Ivermectin-target site (GluCl) associated with field-evolved resistance of head lice recovered from Senegal...... 183 Article 10 : Complexin in Ivermectin resistance in body lice ...... 206
Conclusions et perspectives ...... 245
Références bibliographiques ...... 248
Annexes ...... 251
Article 11 : Multiple Pathogens Including Potential New Species in Tick Vectors in Côte d'Ivoire ...... 252 Article 12 : Molecular identification of microorganisms in chronic wounds, Republic of Guinea (Conakry) ...... 271
Avant-propos
Le format de présentation de cette thèse correspond à une recommandation de la spécialité
Maladies Infectieuses et Microbiologie, à l’intérieur du Master des Sciences de la Vie et de la
Santé qui dépend de l’Ecole Doctorale des Sciences de la Vie de Marseille.
Le candidat est amené à respecter des règles qui lui sont imposées et qui comportent un format de thèse utilisé dans le Nord de l’Europe et qui permet un meilleur rangement que les thèses traditionnelles. Par ailleurs, la partie introduction et bibliographie est remplacée par une revue envoyée dans un journal afin de permettre une évaluation extérieure de la qualité de la revue et de permettre à l’étudiant de commencer le plus tôt possible une bibliographie exhaustive sur le domaine de cette thèse. Par ailleurs, la thèse est présentée sur article publié, accepté ou soumis associé d’un bref commentaire donnant le sens général du travail. Cette forme de présentation a paru plus en adéquation avec les exigences de la compétition internationale et permet de se concentrer sur des travaux qui bénéficieront d’une diffusion internationale.
Professeur Didier RAOULT
1 / 285 Résumé
Les poux de tête (Pediculus humanus capitis) et les poux de corps (P. h. humanus) sont des ectoparasites hématophages obligatoires qui ont coévolué avec les humains depuis des milliers d’années. L'analyse de l’ADN mitochondrial a classé les poux de tête en cinq clades
(A-E) et a placé les poux de corps dans les clades A et D. Actuellement, les poux de corps sont reconnus comme les principaux vecteurs de maladies telles que le typhus épidémique, la fièvre des tranchées et la fièvre récurrente causées par Rickettsia prowazekii, Bartonella quintana, et
Borrelia recurrentis respectivement, alors que les poux de tête quant à eux ne sont considérés que comme vecteurs potentiels.
Dans la première partie de notre thèse, nous avons étudié la diversité génétique des poux anciens et contemporains. Ainsi, l’analyse des poux de tête provenant d'Israël datant de la période romaine a révélé pour la premier fois la présence de clade B au Moyen-Orient, supportant une origine asiatique pour ce clade, suivie par son introduction au Nouveau Monde via les premiers hommes ayant atteint le continent. En outre, nous avons mis en évidence pour la première fois la présence d'un nouveau clade F, démontrant que la diversité génétique des poux est plus élevée qu'on ne le pensait auparavant.
Dans une deuxième partie, nous avons développé une nouvelle technique de PCR en temps réel pour identification moléculaire rapide des clades de poux, puis nous avons étudié les pathogènes qui leurs sont associés. L’analyse d’une large collection de poux provenant de différents pays a mis en évidence la présence de l’ADN de plusieurs bactéries, ce qui renforce l'hypothèse de rôle vectoriel probable des poux de tête. Ainsi, Bartonella quintana a été détectée dans les poux de tête clade E du Mali et Borrelia recurrentis dans des poux de tête de clade A des pygmées de République du Congo. Par ailleurs, d'autres bactéries qui ne sont pas habituellement vectorisées par les poux telles que Coxiella burnetii, Rickettsia aeschlimannii,
Borrelia theileri et de potentielles nouvelles espèces de genre Anaplasma et Ehrlichia ont été
2 / 285 détectées pour la première fois chez les poux. En outre, nous avons mis en évidence une infection massive des poux par plusieurs espèces d’Acinetobacter suggérant que les poux peuvent constituer un hôte préférentiel pour ces bactéries.
Enfin, nous nous sommes intéressés à l'étude des mécanismes de résistance des poux à l’ivermectine. Dans un premier travail, des poux de tête cliniquement résistants à l’ivermectine récoltés au Sénégal ont été analysés en ciblant le gène GluCl, connu pour être impliqué dans la résistance à l’ivermectine. Les résultats obtenus ont mis en évidence la présence de trois mutations faux-sens qui sont retrouvées uniquement chez les poux résistants. Dans un deuxième travail, l’analyse protéomique comparative entre une souche de poux de corps rendue résistante
à l’ivermectine et la souche sensible de référence a révélé que la complexine, une protéine de liaison SNARE jouant un rôle clé dans la régulation de la libération des neurotransmetteurs, a
été la plus significativement down-régulée chez la souche résistante. La suppression de l’expression de la complexine chez des poux sensibles via les ARN interférents a induit leur résistance à l'ivermectine. Ceci représente la première évidence liant cette protéine à la résistance à l’ivermectine et aux insecticides en générale.
Mots-clés : Pediculus humanus ; Phylogénie ; Clades ; Pathogènes ; Ivermectine ; Résistance
3 / 285 Abstract
Head lice, Pediculus capitis humanus, and body lice, Pediculus h. humanus, are obligate blood-sucking parasites that have co-evolved with their human hosts over millions of years.
Head lice are classified into five divergent mitochondrial clades (A, B, C, D, and E) exhibiting some geographic differences, while body lice belong only to clades A and D. Currently, the body lice are the only recognized disease vector of epidemic typhus, trench fever and relapsing fever caused by Rickettsia prowazekii, Bartonella quintana, and Borrelia recurrentis, respectively, while the head lice are so far considered as potential vectors.
In the first part of our thesis we aimed to study the genetic diversity of ancient and contemporary lice. Through the analysis of head lice from Israel of approximately 2,000 years old, we reported for the first time the existence of the clade B in the Middle East, supporting an
Asian origin for this clade followed by its introduction into the New World with the early people. In addition, we highlighted the existence of a sixth mitochondrial clade (Clade F) suggesting that the level of genetic diversity in human lice is higher than previously thought.
Secondary we have developed a new qPCR for a quick molecular identification of all the known clades of lice, which provided to be very useful when analyzing large collections. Then we investigated head-lice-borne associated pathogens by testing several head lice from different countries. The search of head lice-associated microorganisms revealed the presence of the DNA of several bacterial pathogens reinforcing their potential role as vectors of pathogens. Indeed,
Bartonella quintana was detected in head lice of the clade E from Mali. Borrelia recurrentis was detected in head lice of the clade A pygmy individuals from the Republic of the Congo.
Several other bacteria which were not usually associated with lice, such as Coxiella burnetii,
Rickettsia aeschlimannii, Borrelia theileri and potential new species from the Anaplasma and
Ehrlichia were detected for the first time in lice. In addition, we demonstrated a widespread
4 / 285 infection of lice with several species of Acinetobacter, suggesting lice as possible preferential host for these bacteria.
We finally, investigated potential mechanisms underlying resistance to ivermectin in lice, one of the most effective insecticides against lice infestations. For this purpose, samples of wild population of head lice were collected from ivermectin-treated individuals from Dielmo
(Senegal). The clinically confirmed ivermectin resistant individuals were subjected to genetic analysis targeting GluCl gene, the primary target of ivermectin known to be involved in resistance. Using DNA-polymorphism analysis, we have identified, for the first time, the occurrence of three non-synonymous mutations in GluCl gene specific to resistant lice. In a second study, through proteomic analysis of laboratory susceptible and ivermectin-selected resistant body lice, a complexin, a neuronal protein that plays a key role in regulating neurotransmitter release, was shown to be the most significantly down-expressed in ivermectin- resistant lice. Its down-expression by RNA-interference in susceptible lice induced resistance to ivermectin, providing evidence that complexin plays a significant role in regulating ivermectin resistance and represents the first evidence that links complexin to insecticide resistance.
Key words: Pediculus humanus; Phylogeny; Clades; Pathogens; Ivermectin; Resistance
5 / 285 Introduction générale
Les poux hématophages (Phthiraptera : Anoplura) des mammifères sont des ectoparasites spécifiques de leurs hôtes avec qui ils ont coévolué depuis environ 65 millions d’années [1].
L’homme est parasité par deux espèces : Phtirius pubis, communément appelé morpion, appartenant au genre Phtirius qui est partagé avec le gorille (Pt. gorillae), et Pediculus humanus appartenant au genre Pediculus et qui est partagé avec le chimpanzé (P. schaeffi) et les singes du Nouveau Monde (P. mjobergi) [1,2].
P. humanus comprend deux écotypes, le pou de tête (P. h. capitis) et le pou de corps (P. h. humanus), qui sont génétiquement et morphologiquement très similaires, mais occupant chacun une niche écologique différente : les cheveux pour le pou de tête et les vêtements pour le pou de corps [3]. La pédiculose due au pou de tête est très répandue à travers le monde et touche plus particulièrement les enfants, indépendamment de leurs conditions d’hygiène [3].
En revanche, le pou de corps infeste spécialement les populations défavorisées vivant dans des conditions sanitaires extrêmement précaires telles les pauvres, les sans-abris, les prisonniers et les réfugiés de guerre [3].
La diversité génétique des poux du genre Pediculus a été principalement étudiée en utilisant les gènes mitochondriaux (principalement les gènes cytochrome b [cytb] et cytochrome oxydase 1 [cox1]) révélant la présence de cinq clades très divergents (A, D, B, C et E) [4–6].
En plus de cette diversité inter-clade, les poux présentent également une diversité intra-clade, illustrée par plusieurs haplotypes pour chaque clade [2,4]. Les poux de corps appartiennent aux clades A et D, tandis que les poux de tête englobent toute la diversité génétique [5,6]. Le clade
A est le plus répandu sur tous les continents, tandis que les autres clades ont une répartition géographique spécifique [4,7,8]. Le clade D a été retrouvé jusqu’ici en République démocratique du Congo et en Ethiopie [5]. Le clade B a été décrit pour la première fois sur le continent américain, où il est très répandu et diversifié. La présence des membres de ce clade
6 / 285 parmi des poux des momies précolombiennes a amené les chercheurs à inférer une origine américaine pour ce clade, qui a été par la suite redistribué dans le Vieux Monde par les colons ayant regagné l’Europe [8–10]. Ce clade est retrouvé également en Europe, en Australie, en
Afrique du Sud et en Algérie [6,7,11]. Le clade C a été retrouvé en Ethiopie, au Népal, au
Pakistan et au Thaïlande [7,12]. Enfin, le clade E est retrouvé principalement en Afrique de l’ouest à savoir le Senegal et le Mali [13,14].
Les primates et les poux entretiennent une relation qui date d’au moins 25 millions d’années. Les études phylogénétiques des poux basées sur des méthodes de datation moléculaire ont permis de confirmer des événements dans l’histoire évolutive de notre espèce
[1,8]. A juste titre, le P. humanus et le P. schaeffi descendent d’un ancêtre commun datant d’il y a environ 6 millions d’années, ce qui correspond à la datation estimée de la séparation de leurs hôtes respectives, l’Homme et le chimpanzé [1]. Par ailleurs, les poux ont également permis d’élucider des événements dans l'évolution humaine qui ont été incertains des données génétiques et fossiles de l'hôte, tel que l’estimation de la date quand les hommes ont commencé
à porter des vêtements par l’estimation de la séparation de l'ancêtre des P. humanus en P. h. capitis et P. h. humanus il y a environs 83,000 et 170,000 millions d’années [15,16].
Certainement, plus d’études approfondies sur la diversité génétique des poux Pediculus vont permettre d’élucider d'autres événements inconnus ou peu clairs dans l'histoire de l'humanité.
En plus de leur rôle en tant que marqueur de l’évolution, les poux constituent une menace réelle pour l’homme, plus particulièrement le pou de corps qui est le principal vecteur de trois maladies délétères - typhus épidémique, la fièvre des tranchées et la fièvre récurrente causées par Rickettsia prowazekii, Bartonella quintana et Borrelia recurrentis, respectivement - ayant tué des millions de personnes à travers l’histoire de l’humanité [17]. Le pou de corps est
également soupçonné dans la transmission d'un quatrième agent pathogène mortel, Yersinia pestis, l'agent de la peste [18]. Par ailleurs, le rôle du pou de tête en tant que vecteur de maladie
7 / 285 a longtemps été sous-estimé, cependant plusieurs études expérimentales et épidémiologiques laissent fortement entrevoir un rôle vectoriel fort probable pour ce pou [14,19–22]. Néanmoins, sa capacité vectorielle est faible comparée à celle de pou de corps. Ceci est peut-être dû en partie à l’activité phagocytaire de leur système immunitaire, qui est beaucoup plus intense chez le pou de tête comparée au pou de corps, lui permettant ainsi de développer une réponse immunitaire plus efficace vis-à-vis des différents agents pathogènes [22]. A titre d’exemple, la compétence vectorielle du pou de tête a été démontrée pour B. quintana et R. prowazekii dans les conditions expérimentales [21,22]. De plus, l’ADN de plusieurs bactéries telles que B. quintana, B. recurrentis, Y. pestis et Acinetobacter a été mis en évidence dans des poux de tête appartenant à différents clades provenant de plusieurs pays à travers le monde [12,19,20,23,24].
Ces constatations sont particulièrement préoccupantes, parce que ces infestations ne sont contrôlées dans aucun pays, y compris dans les pays les plus riches, et les épidémies continuent
à se produire régulièrement [25]. De plus, la prévalence des poux de corps est sous-estimée dans les pays développés et comme le nombre de personnes sans-abri augmente, l’infestation par les poux ainsi que les maladies transmises sont également à la hausse [26].
En effet, l’accroissement du nombre d’infestations par les poux est principalement dû à l'émergence et à la propagation des populations de poux résistantes aux insecticides conventionnels, en particulier la résistance aux pyréthrinoïdes est devenue majeure, probablement du fait de leur usage intensif dans de nombreux pays [27]. D’autres options thérapeutiques sont actuellement disponibles telle que l’ivermectine, qui semble être une alternative très prometteuse pour lutter contre les poux [28]. Cependant, des cas de résistance potentielle sur le terrain ont été rapportés au Senegal [29]. Par conséquent, la connaissance des mécanismes de résistance est essentielle et permettra de protéger et de prolonger l’activité des insecticides actuels ainsi que pour le développement de nouvelles molécules pour la gestion des résistances déjà décrites. L'accès à la séquence du génome du pou de corps est maintenant
8 / 285 disponible [30] et ouvre de nouvelles perspectives en génomique et génomique fonctionnelle, permettant de mieux comprendre et de mieux caractériser les mécanismes impliqués par des approches moléculaires, transcriptomiques et protéomiques.
Au cours de ce projet de thèse nous avons voulu apporter notre contribution dans le domaine de la recherche sur les poux humains. Dans la première partie de ce manuscrit, nous avons présenté une revue de littérature qui porte sur une synthèse des données actuelles sur les poux, elle s’intéresse notamment aux aspects biologiques, phylogénétiques, épidémiologiques, pathogènes associés, ainsi que les moyens de lutte et l’évolution de la résistance aux insecticides chez les poux. Dans la deuxième partie, nous allons présenter notre travail expérimental, dont les objectifs se répartissent selon trois axes principaux, organisés autour de trois chapitres à savoir (1) la phylogénie et phylogéographie des poux anciens et contemporains, (2) l’épidémiologie des poux et pathogènes associés et (3) l’étude des mécanismes de résistance des poux à l’ivermectine.
9 / 285
Article 1. Revue de literature
Where are we with human lice? A review of the current state of our knowledge
Revue proposée au journal Trends in Parasitology
10 / 285 Where are we with human lice? A review of the current state of our knowledge
Nadia Amanzougaghene1, Florence Fenollar2, Didier Raoult1* and Oleg Mediannikov1*
1Aix-Marseille Univ, IRD, AP-HM, MEPHI, IHU-Méditerranée Infection, Marseille, France 2Aix Marseille Univ, IRD, AP-HM, SSA, VITROME, IHU-Méditerranée Infection, Marseille,
France
*Corresponding authors:
Dr. Oleg MEDIANNIKOV
Phone : +33 (0)4 13 73 24 01 ;Fax : +33 (0)4 13 73 24 02 ;e-mail : [email protected]
Pr. Didier RAOULT
Phone : +33413732401 ; Fax : +33413732402 ; e-mail : [email protected]
Word abstract count: 160
Word text count: 5,543
Tables: 2
Figure: 2
11 / 285 Contents
1. Biology, epidemiology and genomic of lice
2. Phylogeny and phylogeography of lice
2. 1 Phylogeny relationships between head and body lice
2. 2 Louse mitochondrial clades
2. 3 Lice as marker of human evolution
3. Lice-borne bacterial disease
3. 1 Body louse associated pathogens
3. 2 Head louse associated pathogens
4. Control of louse infestations and evolution of insecticide resistance
4. 1 Therapeutic options for pediculosis treatment
4. 2 Insecticide resistance
4. 3 New therapeutic approaches
5. Concluding remarks and Future Perspectives
12 / 285 Abstract Pediculus humanus is an obligate bloodsucking ectoparasite of human that includes two ecotypes, head louse and body louse. These two ecotypes differ slightly in morphology and biology but have distinct ecologies. More importantly, the body louse is the only accepted disease vector that had killed millions. However, the role of head louse as disease vector have been extensively discussed and speculated about in recent years. Since the sequencing of body louse genome, the knowledge about human lice has evolved and has experienced significant advances in our understanding of the phylogeny, taxonomic relationship and the differences in vector competence between head and body lice. Moreover, the transcriptomes of both body and head lice have been completed providing testable hypotheses on important aspects of louse biology. Current lice control strategies have proved unsuccessful, and novel opportunities for pest control strategies are needed. We aim here to give an update of the current knowledge about this fascinating and highly intimate parasite of human.
Keywords: Pediculus humanus, biology, epidemiology, phylogeny, disease-vector, control
13 / 285 1. Biology, epidemiology and genomic of lice
Human lice belong to the insect suborder Anoplura (order: Phthiraptera), more commonly known as the sucking lice and they feed strictly on the blood of their hosts by piercing the skin [1]. More than 530 species have been described and each species parasitizes one or a few closely related host species of placental mammals [1]. Humans are parasitized by two species of lice, Pediculus humanus and Pthirus pubis [2]. Pt. pubis found in the pubic area and known as the pubic or crab louse (not considered further in this review) belongs to the genus Pthirus, which is shared with gorillas (Pt. gorilla) [1,2]. P. humanus is belonged to the genus of Pediculus, which is shared with chimpanzees (P. schaeffi) and New World monkey (P. mjobergi) [1–3].
P. humanus includes two ecotypes, head lice (P. h. capitis) and body lice (P. h. humanus), that are morphologically almost similar, but ecologically distinct and have different feeding patterns [4]. The head louse lives in the scalp region of humans, where the females lay eggs (nits) at the base of hair shafts, and feeds on human blood frequently, every
4–6 hours [4,5]. Body louse lives and lays eggs in clothing, and feeds less frequently but takes more large blood meals than head louse, moreover, lays higher numbers of eggs and develops faster than head louse [5,6]. In addition, body louse is more resistant to environmental conditions, can withstands lower humidity and survives for longer period of time outside the host (48–72 hours)[4].
The head louse infestation is highly prevalent worldwide, particularly in school-aged children, regardless of hygiene conditions, and the transmission occur mainly by head-to-head contact [7]. Adults with poor personal hygiene are also commonly affected [7]. Body louse infestation is less prevalent and is associated with a lack of hygiene, overcrowding conditions and cold weather [8]. For that reason, homeless, jail and refugee populations are predominantly affected [8,9]. Besides to their role as dangerous disease vector (we will
14 / 285 discuss this topic later in this review), louse infestations cause pruritus that may lead to intense irritation [4,7]. Pruritus is often accompanied by excoriations that can become infected secondarily [7]. Post inflammatory pigmentation is also common in chronically infested persons [7].
The recent sequencing and annotation of body louse genome confirmed that P. humanus has the smallest genome of any hemimetabolous insect reported to date and revealed numerous interesting characteristics in the nuclear and mitochondrial genomes, and holds great potential for understanding of coevolution among lice population, symbionts, and pathogens [10]. Its genome about 108 Mb contains 10,773 predicted protein-coding genes and
57 microRNAs [10]. Relative to other insect genomes, the body louse genome contains significantly reduced number of genes associated with environmental sensing and response, including odorant and gustatory receptors and detoxifying enzymes, reflecting its simple life history, that includes humans as their sole host and humans’ blood as the only diet [10,11].
The mitochondrial (mt) genome of body and head lice contains the full complement of
37 genes organized in an unusual architecture of 20 minichromosomes, each minichromosome is 3–4 kb in size and has 1–3 genes and a control region [12]. Notably, this extensively fragmented mt genomes of P. humanus is a fascinating phenomenon and represent the most radical departure to date in bilateral animals from the typical, single circular mt chromosome, and it may be linked to the loss of the gene encoding the mitochondrial single- stranded DNA binding protein involved in mitochondrial genome replication [10,12]. Why and how exactly mt genome became fragmented, however, are still poorly understood.
Body and head lice harbor the same primary endosymbiotic bacteria (Candidatus Riesia pediculicola) that supply the lice with B-vitamins, absent in the human blood [10,13].
Candidatus Riesia pediculicola has small genome that encodes less than 600 genes distributed between a short linear chromosome ( 0.57 Mb) and a circular plasmid ( 8 kb) that includes
∼ ∼
15 / 285 genes required for the synthesis of essential B vitamins [10,14]. The bacterium is housed in specialized structures known as mycetomes located on the ventral side of the midgut and is transmitted from the female louse to its progeny after its migration to the ovaries [15]. It is belongs to the family Enterobacteriaceae within the Candidatus Riesia genus, which is shared with lice that parasitize chimpanzees and gorillas [14,15]. Phylogenetic studies showed that the symbiont has co-evolved with head and body lice and last shared a common ancestor with
P. schaeffi endosymbiont (Candidatus Riesa pediculischaeffi) roughly 5.4 Mya ago [14]. Lice endosymbionts might also support investigations into the evolution of humans [14].
Moreover, aside from to be fundamental for the louse development and survival making them an interesting target for the development of alternative strategy for louse control (we will discuss this topic later in this review), whether this symbiont has any influence on louse behavior or competence as disease vectors is warrant further studies.
2. Phylogeny and phylogeography of lice
2. 1 Phylogeny relationships between head and body lice
Head lice and body lice have distinct ecologies and differ slightly in morphology and biology [4]. Although, for over a century the taxonomic status of these two types of lice has been the subject of debate, they are now considered to be ecotypes of a single species as opposed to separate species [4,5,16]. Despite numerous studies the genetic basis and evolutionary relationships among body and head lice remain obscure as yet.
Until recently, the most predominant opinion was that body louse descended from head louse in nature [4]. Indeed, as the female body louse lays eggs exclusively on the host’s clothing [4,5,8] it was thought that body lice first emerged only recently when modern humans began to wear clothes [5,17], however most of genetics data available today do not support this view. Recently, a new hypothesis for the emergence of body lice has been proposed, suggesting that under certain conditions of low hygiene, a head louse infestation
16 / 285 can transform into a massive infestation provided an opportunity for head louse variants able to ingest a larger blood meal (a characteristic of body lice) to colonize clothing [4,6,9].
Furthermore, several observational studies had also suggested that head lice could become body lice when raised in appropriate conditions [18,19]. Thus, the divergence of the head louse and body louse does not appear to result from as single event, but probably takes place constantly among the two shared louse clades A and D (see below for louse mitochondrial clades), and that this transformation is facilitated by mass infestations [4,6].
Moreover, the transcriptomes analysis of head and body lice resulted in a single gene difference, the PHUM540560 gene which encodes a hypothetical 69-amino acids protein of unknown function, which is thought to be missing in head louse [20]. A subsequent analysis by Drali et al. showed that this gene was also present in the head louse but with a rearranged sequence compared to body louse [21]. Notably, variation in this gene constitutes the only genetic marker identified to date which can distinguish between the two ecotypes once they are removed from their habitat [21]. These findings indicate that head and body lice have almost the same genomic content but are phenotypically different as a result of differential gene expression [4,20].
Indeed, a recent study analyzed alternative splicing using next-generation sequencing data for head and body lice and showed evidence for transcript pool differences between them
[16]. Interestingly, while no functional categories were overrepresented among genes with head louse-specific alternative splicing events, genes containing body louse-specific alternative splicing events were found to be significantly enriched for functional categories in relation with development of the nervous system, salivary gland, trachea, and ovarian follicle cells, as well as regulation of transcription, suggesting that these changes in the developmental program may underpin phenotypic flexibility observed in body lice allowing to them to inhabit clothing [16].
17 / 285 Taken together, these data evidence that the phenotypic shifts associated with the emergence of body lice are likely to be a consequence of regulatory changes, possibly epigenetic in origin, triggered by environmental cues. Such phenotypic modification has been reported to occur in other insects such as in honey bees, termites and migratory locusts [22–
24]. For example, in honey bees the development of queen and worker is strictly controlled by differential feeding of royal jelly and their adult behaviors are accompanied by epigenomic changes [23]. Certainly, further efforts on the genetic studies of head and body lice are needed to link their genetic difference with phenotypic differences. Whole genome sequencing of head lice and comparative genomics combined with transcriptomic, proteomic and epigenomic studies between head and body lice would be useful to better learn about lice and addressing these questions.
2. 2 Louse mitochondrial clades
Genetic studies based on mitochondrial genes (mainly Cytb and Cox1 genes) have inferred to Pediculus lice a robust phylogenetic classification into several divergent clades or haplogroups, exhibiting some geographic differences (Fig. 1) [5,25–27]. Six mitochondrial clades were described (A, D, B, F, C and E) [25–28] (Amanzougaghene et al, unpublished data). In addition to this inter-haplogroup diversity, lice also present intra-haplogroup diversity, illustrated by several distinct haplotypes for each clades (Fig. 1B) [3,26,29,30].
Head lice encompass all clades diversity while body lice belong only to clades A and D [25].
The clade A is the most common and is found around the world, while, clade D is only found in Democratic Republic of Congo (DRC) and in the Republic of Congo (Congo-
Brazzaville), where it was found mainly among pygmy populations [26,27,29,30]. This clade was also reported in lice from Ethiopia and Zimbabwe [29]. Clade B was first described in contemporary lice from the America continent, where it was highly prevalent and diversified
[30–32]. This finding together with its identification in pre-Columbian mummies’ lice have
18 / 285 lead researchers at the beginning to infer an American origin for this clade [33]. However, its recent discover among head lice remains from Israel, dated approximately 2,000 years old, has challenged this assumption and strongly supported a Middle Eastern origin for this clade followed by its introduction into the New World with the early migrants [28]. This clade is also found in Europe, Australia, Algeria, South Africa and Saudi Arabia [26,30,34–36]. Clade
F is the sister group of clade B, was recently described in lice collected from Amerindians of the wayampi community living in isolated and remote village escaped to globalization in
Trois-Sauts (Amanzougaghene et al, unpublished data). P. mjobergi, a New World monkey louse, which is through to be transmitted to monkeys from the first humans reached the
American continent thousands of years ago, is also belonged to this clade [3].
Clade C has been found in Ethiopia, the Republic of Congo and in Asia (Nepal,
Pakistan and Thailand) [25,29,37]. Lastly, clade E consists of head lice from West Africa namely Senegal and Mali where it was highly prevalent [25] and was recently identified in head lice from Nigerian refugees arriving in Algeria and from migrant communities living in
Bobigny, France [38,39]. All these data support the hypothesis that all current human lice travelled with archaic hominids from Africa [6,25].
2. 3 Lice as marker of human evolution
Because human lice are highly host specific and have been evolving in tandem with their primate hosts for a thousand of years, they offer a unique feature to reconstruct human migration and human evolutionary history, thereby complementing the hominin fossil records
[31,40,41]. The studies conducted to date have only begun to tap into this valuable source of information [40]. For instance, phylogenetic analyses of Pediculus lice have confirmed some events in human evolution. For example, chimpanzee lice (P. schaeffi) and human head/body lice last shared a common ancestor roughly six million years ago, which is strikingly similar to the estimated dates for the divergence of their hosts themselves [2,31]. Lice population
19 / 285 show the genetic signature of a recent demographic expansion about 100,000 years ago coinciding with the out-of-Africa expansion of modern human hosts, thereby we can use lice to learn about human migration such as the timing and route of the peopling of the Americas
[30,31]. Reed and colleagues (2004) suggested that tow ancient head louse clades (B and C), whose origin predates modern Homo sapiens, originated on archaic hominins [31]. They found that clade B diverged from clade A between 0.7 and 1.2 MYA, which is similar in age to the ancestor of H. sapiens and H. neanderthalensis, whereas clade C is even more older
(ca. 2 MYA) may have evolved on H. erectus [31]. Because head lice are primarily transmitted horizontally through host-to-host physical contact, this finding suggests that modern humans came into direct contact with archaic hominin species and picked up distinct lineages of head lice [31].
Lice have also elucidated events in human evolutionary history that are missing or uncertain from host fossil or host DNA [35], such as the estimation date of H. sapiens when began wearing clothing by estimating the age of the body louse, which is thought to have evolved only after humans began to wear clothes as this louse live exclusively in clothing
[17]. As recently put forth by Toups and colleagues (2011), based on molecular dating from body louse, suggest that clothing use originated between 83,000 and 170,000 years ago, which is earlier than previously proposed (anywhere from 40 Ka to 3 Ma based on the emergence of eyed needles and the loss of body hair), and suggest that clothing use by H. sapiens likely originated in Africa and may have enabled to them to more readily move into colder climates as they migrated out of Africa and eventually throughout the world [42].
Taken together, all these studies demonstrate how much lice can tell us about mysteries of our evolution. It is likely that further studies of lice and other host-specific parasites of humans, which carries a written record of our past in their DNA, will clarify additional events
20 / 285 in human history while extinct species of hominids are no longer around to give us clues to our origins.
3. Lice-borne bacterial disease
3. 1 Body louse associated pathogens
Body lice are the main vectors of three dangerous human pathogens, namely Rickettsia prowazekii, Borrelia recurrentis and Bartonella quintana (Fig. 2) [8,9]. All these three bacterial pathogens possess genomes that are reduced in size compared to their free-living close relatives [4]. Body lice infection always occurs through blood meals, and the louse remains infected for its entire short life [8]. The transmission of these infections to humans occurs by contamination of bite sites, microlesions of the skin, conjunctivae, and mucous membranes with the faeces or crushed bodies of infected lice [8].
R. prowazekii is the agent of epidemic typhus that has caused substantial health problems during times of war and social disorder [8,43]. Despite the efficacy of antibiotics treatment, the disease remains a major health threat, because it could re-emerge at any moment [9,43]. Indeed, the recovered patients can retain the bacteria for the rest of their life, and under stressful conditions recrudescence may occur as a milder form of typhus, the Brill-
Zinsser disease, that could serve as a source of new epidemics if louse infestation reappears
[9,43]. Furthermore, since R. prowazekii infection can occurs by inhalation, the bacterium is also classified as a biological select agent [43].
B. recurrentis is a spirochete that cause relapsing fever [8]. Historically, massive outbreaks have occurred in Eurasia and Africa, but currently the disease has persisted especially in Ethiopia and neighboring countries, and recently emerged in travelers returning to Europe and North America from endemic regions [44–46].
B. quintana, is most notoriously known as the causative agent of trench fever, but it may also cause a range clinical manifestation including bacillary angiomatosis, endocarditis,
21 / 285 chronic bacteremia and chronic lymphadenopathy [8,9]. The illness was common-place in
France in the 18th century, during Napoleon’s Russian war, and during World Wars I and II
[8,47]. Currently, regarded as re-emerging pathogen in poor countries, as well as in developed countries afflicting a significant portion of homeless populations [9]. For a long time, it was believed that B. quintana was only transmitted by body louse in humans which are thought to be the sole reservoir. Recently, however, B. quintana was found in cats [48] and some human cases have been linked to contact with kittens and cat fleas [49]. Moreover, macaque monkeys and their lice, Pedicinus obtusus, have also been implicated [50,51]. The role of head louse as additional vector was also raised and discussed thereafter.
Recently, there has been an increased emphasis on the role of body lice as vectors for other pathogenic bacteria (Fig. 2). For instance, the combined evidences from both laboratory and epidemiological studies strongly implicate body lice as vector of Yersinia pestis, the causative agent of plague [52,53], and that it may be involved in plague pandemics which correlated better with the epidemiology of louse-borne infections, assumption evidenced through paleomicrobiological studies [54]. Other studies showed that experimentally infected body lice are able to acquire, maintain, and transmit R. conorii (Mediterranean spotted fever,
Indian tick typhus), R. rickettsii (Rocky Mountain spotted fever), which are both transmitted by ticks, and R. typhi (endemic or murine typhus) and R. akari (rickettsial pox), which are usually transmitted by fleas and mites, respectively [55–57]. Furthermore, R. typhi has been isolated from body lice during outbreaks of murine typhus in northern China and India
(Kashmir State) [58,59]. Early field observation in East Africa showed that body lice collected in a place where an epidemic of Q fever occurred three months previously are capable of transmitting Coxiella burnetii to guinea pigs [60,61]. Additionally, it has been shown that, under experimental conditions, it is possible to infect body lice with C. burnetii
[60]. Some other widespread pathogenic bacteria, such as Acinetobacter baumannii, A. lwoffii
22 / 285 and Serratia marcescens have been associated with body lice with the assumption that lice can probably also transmit these agents to humans [62,63].
Together, these studies suggested that body lice obviously may carry a broad spectrum of pathogens bacteria. Moreover, as blood-feeder insect lice are predisposed to uptake, while feeding in their human host, many kinds of other microorganisms including virus and haemoparasites (such as Babesia or Plasmodium). Theoretically it is feasible that lice can transmit any of these agents, being ingested with blood meal if they are capable of surviving in the insect’s midgut [8]. Moreover, the very reduced genome of human lice lacks many genes associated with immune response, environmental sensing and response, including odorant and gustatory receptors and detoxifying enzymes [10,64,65]. Reduced number of defense mechanisms may facilitate the louse infection by different microorganisms.
Therefore, the role of body lice as vectors of these microorganisms is an interesting research topic that is worthy to be addressed in future studies.
3. 2 Head louse associated pathogens
In recent few decades, there has been an increasing recognition of head lice as vector of pathogens, shifting the old-established paradigm which implicated only body lice as disease- vector. Although body lice are currently assumed to be more potent vectors of pathogens, the vector potential of head lice is not yet fully understood. Recently, several studies performed comparative analyses in the immune responses following the bacterial challenge using in vitro-rearing system and showed that body louse compared to head lice exhibited a significantly reduced immune reactions, particularly at the early stage of the immune challenge [65–68]. In one study, Kim and colleagues [66] demonstrated that, as in the case of body lice, head lice can support a persistent load of B. quintana infection for several days following acquisition in a bloodmeal and disseminate viable organisms in their faeces.
However, the rates of proliferation in the gut and numbers of viable B. quintana in the faeces
23 / 285 were significantly higher in body lice compared to head lice [66]. The reduced immune response in body lice could be responsible, in part, for their increased vector competence compared to head lice [65–68], which obviously evolve more efficient immune response to restrict bacterial replication.
Moreover, the fact that body and head lice occupy distinct ecological niches and have distinctly different feeding patterns, are two supplementary factors that might influence pathogens transmission by lice. Body lice live in the clothes where they must deal with host body movements to access to blood, thereby they are suggested to an increased stress compared with head lice [4,8]. The effects of stress on immunity are well known in insect vectors [69] and may cause alterations that may lead to the reduced innate immune response observed in body lice against pathogenic bacteria. Similarly, the body lice are known to take larger blood meal than head lice [4,9]. The uptake of larger infectious blood meals may result in more bacterial entering the louse midgut and hence possibly increased infection.
Under laboratory conditions, head lice can also transmit R. prowazekii. The first evidence was reported by Goldberger and Anderson, who have successfully transmitted typhus to naïve rhesus macaques by subcutaneous inoculation of infected crushed head lice, recovered from patients with epidemic typhus [70]. Their finding was later confirmed by
Murray and Torrey (1975) who showed that head lice feed on infected rabbit with R. prowazekii can be readily infected and disseminate virulent organisms in their faeces, demonstrating that these lice have the potential to be a vector of pathogen under optimal epidemiological conditions [71]. In addition, it has been argued that this louse could also be involved in the transmission and maintenance of this pathogen in nature [72].
In the last few years, the DNA of several pathogenic bacteria are being increasingly detected in head lice in many parts of the world (Fig. 2). Detailed results on the epidemiological studies are reported in Table 1. For instance, B. quintana was the most
24 / 285 frequently detected in head lice belonging to clades A, D, E and C. Furthermore, it is important to note that, except one reported case of its detection in lice collected from school children [73], all other cases occurred on lice infesting deprived populations living in poverty and lacking standard medications (Table 1). An interesting study performed on head lice collected in rural population from Senegal, where several cases of trench fever occur, showed that several head lice were B. quintana positives. These findings together with the absence of body lice from this area for more than thirty years, strongly implicate head lice as the principal acting in maintaining the transmission cycle of B. quintana in the target population
[74].
The B. quintana DNA was also identified in the head louse nits collected from homeless individuals from Marseille indicating the possible vertical transmission of this pathogen [75].
A study performed on lice from persons at various locations in Ethiopia showed that head lice infection by B. quintana was linked to high altitude [76]. Interestingly, all collected head lice from persons living at higher altitudes (>2,121 m) were B. quintana positives whereas at these altitudes, no body lice were infected with B. quintana [76]. This observation raises important questions about the possible relationship of the head lice vectorial capacity and environmental-ecological factors that may drive force underpinning the transmission potential in some geographic area.
B. recurrentis was detected in head lice clade C from patients with louse-borne relapsing fever living in poor region from Ethiopia, and more recently in head lice clade A from hunter-gatherer pygmy individuals in the Republic of the Congo [29,77]. The DNA of Y. pestis was also detected in head lice clade A collected from persons living in a highly plague- endemic area from the eastern Congo [27,52]. Other human pathogenic bacteria which are not usually associated with lice transmission, such as C. burnetii and R. aeschlimannii, as well as
25 / 285 the DNA of B. theileri and potential new species from the Anaplasma and Ehrlichia genera, of unknown human pathogenicity, were also detected in head lice [25,29].
Most likely the infection of head lice with all the pathogenic bacteria cited above occurs in more vulnerable and deprived populations living in poverty and lack of hygienic, the same conditions that usually lead to body lice proliferation and emergence of louse-borne diseases
[8,9]. This view is also supported by the fact that most of studies conducted in head lice infesting school children and population living in more hygienic conditions were failed to detect theses pathogenic bacteria [37,78–80].
However, head lice infection with Acinetobacter is an exception, as several species have been detected with high prevalence in head lice as well as in body lice everywhere they have been sought, reflecting the ubiquitous occurrence of these bacterial species [29,37,78,79].
This widespread infection of human lice suggests that they could be a preferential host for these bacteria. However, it remains to be determined whether these Acinetobacter strains present in lice are the same as those that are responsible for human infections [29,78]. More attention is now paid to extra-hospital reservoirs of these opportunistic bacteria and their potential involvement in emerging human community-acquired infections, as pan drug resistant strains are increasingly being identified worldwide [81].
Based on the combined evidence of both epidemiological and laboratory studies, we believe that head lice can transmit disease to their human host under favorable epidemiological conditions, although its vectorial capacity is lower compared to body lice.
Therefore, given the scale of head louse infestations around the world and, the emergence and spread of insecticides resistance, this pest are warrants more attention as a serious public health problem.
4. Control of lice infestations and evolution of insecticide resistance
4. 1 Therapeutic options for pediculosis treatment
26 / 285 There are numerous treatment options for pediculosis including chemical insecticides, topically applied physically acting agents (Table 2), herbal formulations and mechanical methods (combs and heating devices) [44,82,83]. Although, the use of chemical insecticides, with a neurotoxic mode of action, is still the method of choice and the most extensively used approach. They are either organophosphates (malathion), organochloride (lindane), carbamates (carbaryl), pyrethrins (extract of chrysanthemum) or pyrethroids (synthetic derivates of pyrethrins such as permethrin, phenothrin or deltamethrin) (Table 2) [44,84,85].
Among them, malathion and permethrin remain the most common used pediculicides since they became available in 1971 and 1992, respectively [84,85].
Unfortunately, the extensive use of these insecticides has led to the emergence and spread of resistant louse populations all over the world [84,85]. This has prompted research into the development of compounds with other modes of action. Ivermectin and spinosad appear to be the most promising new treatments, raised their interest for owing their novel neurotoxic modes of action, low mammalian toxicity and little cross-resistance with commonly used groups of pediculicides [44,86–88]. Furthermore, ivermectin is the only drug currently used for oral treatment, and its highly effectiveness was clinically approved for both head and body lice treatment [88,89], although empirically noted ivermectin resistance in the field began to be reported [90].
There is also increasing interest in the use of natural products such as pediculicides based on plant-derived essential oils (eucalyptus oil and tea tree oil) or with a pure physical mode of action, such as dimeticone and benzyl alcohol [91–93], but there has been little interest in evaluating them clinically for effectiveness, even though some of them are already marketed.
4. 2 Insecticide resistance
27 / 285 Insecticide resistance resulting in treatment failure is believed to be a major factor in the increasing number of infestations by lice [44,85]. Resistance is an acquired trait that an insect pest develops over time through selective pressure created by prolonged or inadequate use of insecticides [85]. The recently sequenced body louse genome offers unique opportunity for addressing fundamental questions relating to the molecular mechanisms driving the insecticides resistances, that is essential to maximize the active lifespan of existing insecticides, and to accelerate the development of new strategies and tools towards fighting louse infestations [10,86].
Possible mechanisms of resistance include knockdown-resistance in the case of permethrin, which is the result of three-point mutations (M815I, T917I and L920) within the
α-subunit gene of the voltage-gated sodium channel, and enhanced activity of a carboxylesterase enzyme was found to be responsible for the malathion resistance [94,95].
Recently, researcher of our team reported the first field-evolved ivermectin-resistance in head lice occurring in rural populations of Senegal [90]. Genetic analysis of these lice targeting
GluCl gene, the primary target-site of ivermectin known to be involved in resistance in arthropods and nematodes, revealed the presence of three relevant non-synonymous mutations
(A251V, H272R and S46P) that may be responsible for the treatment failure
(Amanzougaghene et al, unpublished data). Furthermore, in another study we found that through proteomic analysis of laboratory susceptible and ivermectin-selected resistant body lice, a complexin, a neuronal protein that plays a key role in regulating neurotransmitter release, was shown to be the most significantly down-expressed protein in ivermectin- resistant lice. Its down-expression by RNA-interference in susceptible lice induced resistance to ivermectin, providing evidence that complexin plays a significant role in regulating ivermectin resistance, linking for the first time complexin to insecticide resistance
(Amanzougaghene et al, unpublished data).
28 / 285 4. 3 New therapeutic approaches
Symbiotic therapy. This approach incited great interest for potential applications in public health entomology [96] and has the advantage of targeting the bacteria, which are sensitive to the action of antibiotics [96]. Since the primary symbiont of human lice,
Candidatus Riesia pediculicola, is essential to the metabolism of the host, due to its capacity to synthesize B-group vitamins, the deleterious action exerted by the antibiotic treatment should reflect on the host [10,82]. A first case report showed that the antibiotic treatment
(trimethoprim and sulfamethoxazole), performed to combat a respiratory infection in a 12- year-old girl resulted as a side effect in the death of head lice [97]. Subsequently, studies conducted by Sangaré et al, demonstrated the effectiveness of this therapy under laboratory conditions and showed that antibiotics (such as doxycycline, erythromycin, rifampicin and azithromycin) kills human lice through its direct activity on their Riesia symbiont [98].
Moreover, the combination of this antibiotic with ivermectin provided to be highly effectiveness compared with ivermectin alone in treating and preventing body lice and could be used to achieve complete eradication of lice and potentially delay the emergence of ivermectin resistance [99]. This approach is promising therapy but has yet to be evaluated in field studies.
RNAi-based insecticides. RNA interference (RNAi) technology is a promising environmental friendly method to control insects by double-stranded RNA (dsRNA) or small interfering RNA (siRNA) triggered post-transcriptional gene silencing [100]. RNAi can induce mortality, create beneficial phenotypes for insect control, and prevent insecticide resistance in insect pests [101,102]. Currently, the potential of RNAi for future management of pest insects is widely recognized and holds great promise [101,103]. Therefore, it is exciting to consider its role in lice control as alternative to chemical insecticides by suppressing essential genes leading to RNAi-induced mortality [104]. Two lines of evidence
29 / 285 support its potential use as control strategy in lice. First, body louse genome analysis has been shown to contain the genes necessary for RNAi machinery [11]. Subsequently, studies have reported that the injection of dsRNA can effectively suppress target genes in both body and head lice [95,105]. A second advantage of using this approach in lice is lack of gene redundancy within its small genome. Thus, a smaller set of genes can be tested to determine which one is critical for a given biological process [11]. A priority for the future should be focused in identifying efficient RNAi target genes and subsequently exploring method of delivery using field feasible applications [11]. Lastly, even if it is technically feasible, it remains to be determined if such approaches would be approved by regulatory agencies and if an economically viable strategy can be developed to employ RNAi strategies to lice control
[11].
5. Concluding remarks and future perspectives
In the 21st century, human lice infestation remains prevalent worldwide. Surprising and new insights into the biology, epidemiology, and the evolutionary history of lice, their bacterial disease agents and control strategies still stimulated a renewal of interest in this bloodsucking insect. Over the past few years, knowledge about lice has evolved, with the sequencing of body louse genome and the accomplishment of both body and head lice transcriptomes. However, the functional genomics of critical aspects of lice biology is still in its infancy and many aspects are not completely understood and remain to be uncovered.
Further efforts on the genetic studies of head and body lice are needed. Whole genome sequencing of head lice belonging to different clades with integration of technologies developed to observe global changes in transcription and translation, as well as computational tools will accelerate the addressing of important biological questions, identification and exploitation of new target genes in this insect vector, insecticide discovery, including mechanisms of resistance to existing treatments, as well as to develop novel therapies.
30 / 285 Although, our knowledge increases concerning the vector competence of head lice, there is still a need to explore factors that can influence vector competence between body and hand lice, such as the influence of the immune responses and microbiota (especially the role of endosymbiotic bacteria), once addressed, will provide important insights into to effective strategies of lice control and for preventing re-emerging diseases. Finally, because P. humanus is probably one of the oldest and highly intimate human parasites which carries a written record of our past in its DNA, integrating phylogenomic and population genomic patterns in lice will provides a more complete picture of parasite evolution and will clarify additional events in our evolutionary history.
31 / 285 Acknowledgments
This study was supported by the Fondation Méditerranée Infection and the French National
Research Agency under the “Investissements d’avenir” program, reference ANR-10-IAHU-
03.
32 / 285 References
1 Durde , L.A. a d Musser, G.G. The su ki g li e I se ta, A oplura of the orld : a taxonomic checklist with records of mammalian hosts and geographical distributions. Bulletin of the AMNH ; o. . Sucking lice and hosts at
33 / 285 24 Simpson, S.J. et al. (2001) Gregarious behavior in desert locusts is evoked by touching their back legs. Proc. Natl. Acad. Sci. U. S. A. 98, 3895–3897 25 Amanzougaghene, N. et al. (2017) Detection of bacterial pathogens including potential new species in human head lice from Mali. PLOS ONE 12, e0184621 26 Ashfaq, M. et al. (2015) High diversity and rapid diversification in the head louse, Pediculus humanus (Pediculidae: Phthiraptera). Sci. Rep. 5, 27 Drali, R. et al. (2015) A New Clade of African Body and Head Lice Infected by Bartonella quintana and Yersinia pestis-Democratic Republic of the Congo. Am. J. Trop. Med. Hyg. 93, 990– 993 28 Amanzougaghene, N. et al. (2016) High Ancient Genetic Diversity of Human Lice, Pediculus humanus, from Israel Reveals New Insights into the Origin of Clade B Lice. PloS One 11, e0164659 29 Amanzougaghene, N. et al. (2016) Head Lice of Pygmies Reveal the Presence of Relapsing Fever Borreliae in the Republic of Congo. PLoS Negl. Trop. Dis. 10, e0005142 30 Ascunce, M.S. et al. (2013) Mitochondrial diversity in human head louse populations across the Americas. Am. J. Phys. Anthropol. 152, 118–129 31 Reed, D.L. et al. (2004) Genetic Analysis of Lice Supports Direct Contact between Modern and Archaic Humans. PLOS Biol. 2, e340 32 Light, J.E. et al. (2008) Geographic distributions and origins of human head lice (Pediculus humanus capitis) based on mitochondrial data. J. Parasitol. 94, 1275–1281 33 Boutellis, A. et al. (2013) Evidence of sympatry of clade a and clade B head lice in a pre- Columbian Chilean mummy from Camarones. PloS One 8, e76818 34 Boutellis, A. et al. (2015) Evidence that clade A and clade B head lice live in sympatry and recombine in Algeria. Med. Vet. Entomol. 29, 94–98 35 Raoult, D. et al. (2008) Molecular identification of lice from pre-Columbian mummies. J. Infect. Dis. 197, 535–543 36 Al-Shahrani, S.A. et al. (2017) Genetic diversity of the human head lice, Pediculus humanus capitis, among primary school girls in Saudi Arabia, with reference to their prevalence. Parasitol. Res. 116, 2637–2643 37 Sunantaraporn, S. et al. (2015) Molecular survey of the head louse Pediculus humanus capitis in Thailand and its potential role for transmitting Acinetobacter spp. Parasit. Vectors 8, 127 38 Candy, K. et al. (2018) Molecular Survey of Head and Body Lice, Pediculus humanus, in France. Vector Borne Zoonotic Dis. Larchmt. N DOI: 10.1089/vbz.2017.2206 39 Louni, M. et al. (2018) Detection of bacterial pathogens in clade E head lice collected from Niger’s refugees i Algeria. Parasit. Vectors 11, 348 40 Perry, G.H. (2014) Parasites and human evolution. Evol. Anthropol. Issues News Rev. 23, 218– 228 41 Boutellis, A. et al. (2014) The origin and distribution of human lice in the world. Infect. Genet. Evol. J. Mol. Epidemiol. Evol. Genet. Infect. Dis. 23, 209–217 42 Toups, M.A. et al. (2011) Origin of Clothing Lice Indicates Early Clothing Use by Anatomically Modern Humans in Africa. Mol. Biol. Evol. 28, 29–32 43 Bechah, Y. et al. (2008) Epidemic typhus. Lancet Infect. Dis. 8, 417–426 44 Bonilla, D.L. et al. (2013) The biology and taxonomy of head and body lice--implications for louse-borne disease prevention. PLoS Pathog. 9, e1003724 45 Antinori, S. et al. (2016) Louse-borne relapsing fever among East African refugees in Europe. Travel Med. Infect. Dis. 14, 110–114 46 Antinori, S. et al. (2016) Louse-Borne Relapsing Fever (Borrelia recurrentis) in a Somali Refugee Arriving in Italy: A Re-emerging Infection in Europe? PLoS Negl. Trop. Dis. 10, e0004522 47 Raoult, D. et al. (2006) Evidence for louse-tra s itted diseases i soldiers of Napoleo ’s Gra d Army in Vilnius. J. Infect. Dis. 193, 112–120 48 Drancourt, M. et al. (1996) Bartonella (Rochalimaea) quintana infection in a seronegative hemodialyzed patient. J. Clin. Microbiol. 34, 1158–1160
34 / 285 49 Raoult, D. et al. (1994) Bartonella (Rochalimaea) quintana isolation in patient with chronic adenopathy, lymphopenia, and a cat. Lancet Lond. Engl. 343, 977 50 Li, H. et al. (2013) Transmission and Maintenance Cycle of Bartonella quintana among Rhesus Macaques, China. Emerg. Infect. Dis. 19, 297–300 51 Sato, S. et al. (2015) Japanese Macaques (Macaca fuscata) as Natural Reservoir of Bartonella quintana. Emerg. Infect. Dis. 21, 2168–2170 52 Piarroux, R. et al. (2013) Plague epidemics and lice, Democratic Republic of the Congo. Emerg. Infect. Dis. 19, 505–506 53 Houhamdi, L. et al. (2006) Experimental model to evaluate the human body louse as a vector of plague. J. Infect. Dis. 194, 1589–1596 54 Raoult, D. (2016) A Personal View of How Paleomicrobiology Aids Our Understanding of the Role of Lice in Plague Pandemics. Microbiol. Spectr. 4, 55 Houhamdi, L. et al. (2003) An experimental model of human body louse infection with Rickettsia typhi. Ann. N. Y. Acad. Sci. 990, 617–627 56 Houhamdi, L. and Raoult, D. (2006) Experimentally infected human body lice (pediculus humanus humanus) as vectors of Rickettsia rickettsii and Rickettsia conorii in a rabbit model. Am. J. Trop. Med. Hyg. 74, 521–525 57 Weyer, F. (1952) The behavior of Rickettsia akari in the body louse after artificial infection. Am. J. Trop. Med. Hyg. 1, 809–820 58 Kalra, S.L. and Rao, K.N.A. (1951) Typhus Fevers In Kashmir State. Part II. Murine Typhus. Indian J. Med. Res. 39, 297–302 59 Liu, W.T. (1944) Studies on the Murine Origin of Typhus Epidemics in North China. 3. Isolation of Murine Typhus Rickettsia from Rats, Rat-Fleas and Body-Lice of Patients during an Epidemic in a Poor House. Chin. Med. J. (Engl.) 62, 119–39 60 Babudieri, B. (1959) Q fever: A zoonosis. Adv Vet Sci 5, 82–182 61 Giroud, P. a d Jadi , J. I fe tio late te et o ser atio de Ri kettsia ur etii hez l’ho e, le rôle du pou. Bull Soc. Pathol. exotique 47, 764–765 62 Houhamdi, L. and Raoult, D. (2006) Experimental infection of human body lice with Acinetobacter baumannii. Am. J. Trop. Med. Hyg. 74, 526–531 63 La Scola, B. et al. (2001) Detection and Culture of Bartonella quintana, Serratia marcescens, and Acinetobacter spp. from Decontaminated Human Body Lice. J. Clin. Microbiol. 39, 1707– 1709 64 Lee, S.H. et al. (2010) Decreased detoxification genes and genome size make the human body louse an efficient model to study xenobiotic metabolism. Insect Mol. Biol. 19, 599–615 65 Previte, D. et al. (2014) Differential gene expression in laboratory strains of human head and body lice when challenged with Bartonella quintana, a pathogenic bacterium. Insect Mol. Biol. 23, 244–254 66 Kim, J.H. et al. (2017) Comparison of the proliferation and excretion of Bartonella quintana between body and head lice following oral challenge. Insect Mol. Biol. 26, 266–276 67 Kim, J.H. et al. (2011) Comparison of the humoral and cellular immune responses between body and head lice following bacterial challenge. Insect Biochem. Mol. Biol. 41, 332–339 68 Kim, J.H. et al. (2012) Comparison of the immune response in alimentary tract tissues from body versus head lice following Escherichia coli oral infection. J. Asia-Pac. Entomol. 15, 409–412 69 Muturi, E.J. et al. (2011) Larval environmental stress alters Aedes aegypti competence for Sindbis virus. Trop. Med. Int. Health TM IH 16, 955–964 70 Goldberger, J. and Anderson, J.F. (1912) The Transmission of Typhus Fever, with Especial Reference to Transmission by the Head Louse (Pediculus capitis). Public Health Rep. 1896-1970 27, 297–307 71 Murray, E.S. and Torrey, S.B. (1975) Virulence of Rickettsia prowazeki for head lice. Ann. N. Y. Acad. Sci. 266, 25–34 72 Robinson, D. et al. (2003) Potential role of head lice, Pediculus humanus capitis, as vectors of Rickettsia prowazekii. Parasitol. Res. 90, 209–211
35 / 285 73 Eremeeva, M.E. et al. (2017) Molecular Markers of Pesticide Resistance and Pathogens in Human Head Lice (Phthiraptera: Pediculidae) From Rural Georgia, USA. J. Med. Entomol. 54, 1067–1072 74 Diatta, G. et al. (2014) Prevalence of Bartonella quintana in patients with fever and head lice from rural areas of Sine-Saloum, Senegal. Am. J. Trop. Med. Hyg. 91, 291–293 75 Angelakis, E. et al. (2011) Bartonella quintana in head louse nits. FEMS Immunol. Med. Microbiol. 62, 244–246 76 Angelakis, E. et al. (2011) Altitude-dependent Bartonella quintana genotype C in head lice, Ethiopia. Emerg. Infect. Dis. 17, 2357–2359 77 Boutellis, A. et al. (2013) Borrelia recurrentis in head lice, Ethiopia. Emerg. Infect. Dis. 19, 796– 798 78 Candy, K. et al. (2018) Molecular Survey of Head and Body Lice, Pediculus humanus, in France. Vector-Borne Zoonotic Dis. DOI: 10.1089/vbz.2017.2206 79 Bouvresse, S. et al. (2011) No evidence of Bartonella quintana but detection of Acinetobacter baumannii in head lice from elementary schoolchildren in Paris. Comp. Immunol. Microbiol. Infect. Dis. 34, 475–477 80 Fournier, P.-E. et al. (2002) Human pathogens in body and head lice. Emerg. Infect. Dis. 8, 1515–1518 81 Eveillard, M. et al. (2013) Reservoirs of Acinetobacter baumannii outside the hospital and potential involvement in emerging human community-acquired infections. Int. J. Infect. Dis. IJID Off. Publ. Int. Soc. Infect. Dis. 17, e802-805 82 Sangaré, A.K. et al. (2016) Management and Treatment of Human Lice. BioMed Res. Int. 2016, 83 Feldmeier, H. (2014) Treatment of pediculosis capitis: a critical appraisal of the current literature. Am. J. Clin. Dermatol. 15, 401–412 84 Clark, J.M. et al. (2013) Human lice: Past, present and future control. Pestic. Biochem. Physiol. 3, 162–171 85 Durand, R. et al. (2012) Insecticide resistance in head lice: clinical, parasitological and genetic aspects. Clin. Microbiol. Infect. Off. Publ. Eur. Soc. Clin. Microbiol. Infect. Dis. 18, 338–344 86 Clark, J.M. et al. (2015) Utilization of the human louse genome to study insecticide resistance and innate immune response. Pestic. Biochem. Physiol. 120, 125–132 87 Aditya, S. and Rattan, A. (2012) Spinosad: An effective and safe pediculicide. Indian Dermatol. Online J. 3, 213–214 88 Chosidow, O. et al. (2010) Oral ivermectin versus malathion lotion for difficult-to-treat head lice. N. Engl. J. Med. 362, 896–905 89 Foucault, C. et al. (2006) Oral ivermectin in the treatment of body lice. J. Infect. Dis. 193, 474– 476 90 Diatta, G. et al. (2016) Head lice probably resistant to ivermectin recovered from two rural girls in Dielmo, a village in Sine-Saloum, Senegal. Int. J. Antimicrob. Agents 47, 501–502 91 Candy, K. et al. (2018) In vitro efficacy of five essential oils against Pediculus humanus capitis. Parasitol. Res. 117, 603–609 92 Meinking, T.L. et al. (2010) The clinical trials supporting benzyl alcohol lotion 5% (Ulesfia): a safe and effective topical treatment for head lice (pediculosis humanus capitis). Pediatr. Dermatol. 27, 19–24 93 Burgess, I.F. et al. (2013) Single application of 4% dimeticone liquid gel versus two applications of 1% permethrin creme rinse for treatment of head louse infestation: a randomised controlled trial. BMC Dermatol. 13, 5 94 Clark, J.M. (2009) Determination, mechanism and monitoring of knockdown resistance in permethrin-resistant human head lice, Pediculus humanus capitis. J. Asia-Pac. Entomol. 12, 1–7 95 Kwon, D.H. et al. (2014) Identification and characterization of an esterase involved in malathion resistance in the head louse Pediculus humanus capitis. Pestic. Biochem. Physiol. 112, 13–18 96 Sassera, D. et al. (2013) Microbial symbiosis and the control of vector-borne pathogens in tsetse flies, human lice, and triatomine bugs. Pathog. Glob. Health 107, 285–292
36 / 285 97 Shashindran, C.H. et al. (1978) Oral therapy of pediculosis capitis with cotrimoxazole. Br. J. Dermatol. 98, 699–700 98 Sangaré, A.K. et al. (2015) Doxycycline kills human lice through its activity on their bacterial symbiont. Int. J. Antimicrob. Agents 45, 675–676 99 Sangaré, A.K. et al. (2016) Synergistic activity of antibiotics combined with ivermectin to kill body lice. Int. J. Antimicrob. Agents 47, 217–223 100 Das, S. et al. (2015) Chitosan, Carbon Quantum Dot, and Silica Nanoparticle Mediated dsRNA Delivery for Gene Silencing in Aedes aegypti: A Comparative Analysis. ACS Appl. Mater. Interfaces 7, 19530–19535 101 Airs, P.M. and Bartholomay, L.C. (2017) RNA Interference for Mosquito and Mosquito-Borne Disease Control. Insects 8, 102 Gordon, K.H.J. and Waterhouse, P.M. (2007) RNAi for insect-proof plants. Nat. Biotechnol. 25, 1231–1232 103 Baum, J.A. et al. (2007) Control of coleopteran insect pests through RNA interference. Nat. Biotechnol. 25, 1322–1326 104 Scott, J.G. et al. (2013) Towards the elements of successful insect RNAi. J. Insect Physiol. 59, 105 Yoon, K.S. et al. (2011) Brief exposures of human body lice to sub-lethal amounts of ivermectin over transcribes detoxification genes involved in tolerance. Insect Mol. Biol. 20, 687–699 106 Sasaki, T. et al. (2006) First molecular evidence of Bartonella quintana in Pediculus humanus capitis (Phthiraptera: Pediculidae), collected from Nepalese children. J. Med. Entomol. 43, 110– 112 107 Bonilla, D.L. et al. (2009) Bartonella quintana in body lice and head lice from homeless persons, San Francisco, California, USA. Emerg. Infect. Dis. 15, 912–915 108 Bonilla, D.L. et al. (2014) Risk factors for human lice and bartonellosis among the homeless, San Francisco, California, USA. Emerg. Infect. Dis. 20, 1645–1651 109 Cutler, S. et al. (2012) Bartonella quintana in Ethiopian lice. Comp. Immunol. Microbiol. Infect. Dis. 35, 17–21 110 Boutellis, A. et al. (2012) Bartonella quintana in head lice from Sénégal. Vector Borne Zoonotic Dis. Larchmt. N 12, 564–567 111 Sangaré, A.K. et al. (2014) Detection of Bartonella quintana in African body and head lice. Am. J. Trop. Med. Hyg. 91, 294–301 112 Kempf, M. et al. (2012) Detection of Acinetobacter baumannii in human head and body lice from Ethiopia and identification of new genotypes. Int. J. Infect. Dis. IJID Off. Publ. Int. Soc. Infect. Dis. 16, e680-683 113 Mana, N. et al. (2017) Human head lice and pubic lice reveal the presence of several Acinetobacter species in Algiers, Algeria. Comp. Immunol. Microbiol. Infect. Dis. 53, 33–39 114 Burgess, I.F. et al. (2012) 1,2-Octanediol, a novel surfactant, for treating head louse infestation: identification of activity, formulation, and randomised, controlled trials. PloS One 7, e35419 115 Burgess, I.F. and Silverston, P. (2015) Head lice. BMJ Clin. Evid. 2015,
37 / 285
Fig 1. Phylogeography of body and head lice haplogroups. A) Neighbor-joining tree based on Cytb haplotypes. B) Median Joining Network representing the relationships between different haplotypes. Pie colors and sizes in circles represent the continents and the number of their sequence for a haplotype. C) Maps of the world showing the distribution of louse clades.
38 / 285
Fig 2. Body and head lice associated bacterial pathogens.
39 / 285 Table 1. Bacteria species found in head lice using molecular methods
Pathogen Lice tested Louse- Collection Population Country Co-occurrence Comment Reference (% positive) clade year of body louse B. quintana 21 (9.5%) _ 2002 Homeless and school Nepal Yes All lice collected from school [106] children children were B. quintana negatives 15 pools _ 2007-2008 Homeless persons California- Not all _ [107] (20%) USA 16 pools _ 2008-2010, Homeless persons California- Not all _ [108] (37.5%) 2012 USA 3 pools nits _ _ Homeless persons France No Attempt to cultivate B. quintana [75] (100%) from these nits was failed 35 (17.1%) A and 2010 Persons living in a highly Congo RDC Yes _ [27,52] D plague-endemic area 35 (2.85%) - 2011 Patients with Ethiopia Yes _ [77] louse-borne relapsing fever
40 / 285 65 pools _ 2010 Street beggars (in poorer Ethiopia _ More pools of head lice were found [109] (9.2%) regions) positive than body lice 271 (7%) C _ Persons living at Ethiopia Not all B. quintana in head lice was [76] locations at different positively linked to altitude (>2,121 altitudes m). At this altitude all body lice were B. quintana negatives 274 (6.93%) A and 2010-2011 Persons living in poor Senegal No _ [110] E conditions 381 (0.52) A 2011 Rural community Senegal _ _ [111]
148 (1.3%) A 2011 Rural villagers living in Senegal No Co-occurrence of trench fever cases [74] poor conditions with absence of body lice for more than 30 years in the studied area 75 (2.66) A 2010–2011 Rural community with Madagascar _ _ [111] low income 600 (0.5%) E 2013 Rural villagers living in Mali No Apparently healthy individuals, low [25] poor conditions socioeconomic level 168 (10.3%) 2013-2015 School children Georgia-USA No The kdr-permethrin resistance [73] (T917I mutation) was detected in 99.9% of 168 lice tested B. recurrentis 35 (22.8%) _ 2011 Patients with louse-borne Ethiopia Not all 4 of 5 patients were co-infested [77] relapsing fever with body louse 630 (1.6%) A 2015 Pygmy populations, Republic of No _ [29] living in poor conditions Congo Y. pestis 35 (2.86%) A 2010 Persons living in a highly Congo RDC Yes _ [27,52] plague-endemic area B. theileri 630 (0.16%) A 2015 Pygmy populations, Republic of No _ [29] living in poor conditions Congo C. burnetii 600 (1.16%) E 2013 Rural villagers living in Mali No MST genotype 35 and new [25] poor conditions genotype (genotype 59) 37 (8.10%) E 2016 Niger’s refugees arriving Algeria No _ [39] in Algeria, living in poor conditions R. 600 (0.6%) E 2013 Rural villagers living in Mali No Apparently healthy individuals, low [25] aeschlimannii poor conditions socioeconomic level Anaplasma 600 (0.3%) E 2013 Rural villagers living in Mali No potential new species [25] poor conditions Ehrlichia 600 (2.3%) E 2013 Rural villagers living in Mali No E. aff. mineirensis and potential [25]
41 / 285 poor conditions new species Acinetobacter 288 (33%) _ 2008-2009 School children France _ _ [79] 115 (47%) A and _ Healthy individuals Ethiopia Yes 13 of 23 lice sequenced were A. [112] C baumannii 275 (3.62%) A and 2013-2014 School children Thailand No Species: A. baumannii, A. [37] C radioresistens and A. schindleri 630 (31.1%) A, C 2015 Pygmy populations, Republic of No Species: A. junii, A. ursingii, A. [29] and D living in poor conditions Congo baumannii, A. johnsonii, A. schindleri, A. lwoffii, A. nosocomialis and A. towneri 52 (80.8%) _ 2013-2015 School children Georgia-USA No _ [73] 235 (11.5%) A, B 2015-2016 Middle-class suburban France No A. baumannii, A. calcoaceticus, A. [78] and E families nosocomialis, A. junii and 2 potential new species (Candidatus Acinetobacter Bobigny-1 and 2) 64 (27%) A and 2014 School children Algeria No A. baumannii, A. johnsonii and A. [113] B variabilis 37 (46.94%) E 2016 Niger’s refugees arriving Algeria No A. baumannii [39] in Algeria Moraxellaceae 630 (0.95%) A 2015 Pygmy populations, Republic of No New species [29] living in poor conditions Congo Table 2. Main Therapeutic options for pediculosis treatment
Pediculicide Class Mechanism of action Adulticide Documented Documented References / Ovicidal adverse health resistance in activities effect lice
Pediculicide with a Neurotoxic Mode of Action
DDT, Organochloride Opening of sodium ion channels Yes/Yes Toxic Yes [44,85] dichlorodiphenyltrichloroethane in neurons
Lindane Organochloride Inhibition of c-aminobutyric Yes/Yes Toxic Yes [44,85] acid- gated chloride channel
Natural pyrethrins Chrysanthemum extract Delayed repolarization of Yes/No Minor Yes [44] voltage-gated
Permethrin, synthetic pyrethrin (+)-3-phenoxybenzyl 3-(2,2- The same as natural pyrethrins Yes/No Minor Yes [44,84,85]
42 / 285 dichlorovinyl)-2,2, dimethyl cyclopropan carboxylate
Malathion Organophosphate Irreversible inhibition of Yes/No Minor Yes [44,85,95] acetylcholinesterase
Carbaryl Carbamate Irreversible inhibition of Yes/No Moderate to Yes [44,85] acetylcholinesterase very toxic
Ivermectin* Macrocyclic lactone Binding to glutamate-gated Yes/No None to Yes [84,88,90] chloride ion channels minimal
Spinosad Macrocyclic lactone Overstimulates nerve cells by Yes/Yes Minor No [83,87] acting like acetylcholine Pediculicide with physical mode of action
Dimeticone Synthetic silicone oils Work by occlusion Yes/Yes Low No [83,85,93]
Isopropyl myristate Ester Work by occlusion or by Only head Minimal No [83] dissolving cuticle wax lice tested 1,2-octanediol Detergent Dehydration by reducing the Yes/No Minimal No [83,114,115] ability of louse to prevent water loss through the cuticle
Benzyl alcohol Aromatic alcohol Asphyxiates lice by ‘‘stunning’’ Yes/No Minimal No [92] the spiracles open
*Ivermectin is the only pediculicide used on topical and oral administrations whereas the other pediculicides are only available through topical applications 43 / 285
Chapitre I :
Phylogénie et phylogéographie des poux anciens et contemporains
44 / 285 Préambule
Dans sa conquête du monde l’Homo sapiens a toujours été en symbiose avec ses poux,
Pediculus, c’est pourquoi ils sont considérés comme de véritables marqueurs de divergence et de migration de notre espèce au fil du temps [1,4]. Les nombreuses études phylogénétiques basées sur les analyses des gènes mitochondriaux ont montré que les poux humains sont distribués à l’intérieur de cinq divergents clades (A, D, B, C et D) qui, à l’exception de clade A qui possède une distribution mondiale, ont une répartition géographique spécifique [4–6]. Dans ce premier chapitre nous rapportons des résultats phylogénétiques que nous avons obtenu grâce
à l’analyse de poux humains rares et précieux.
Un premier travail d'analyse des poux de tête provenant d'Israël de la période romaine, a révélé pour la premier fois la présence de clade B dans cette région, suggérant une origine non- américain pour ce clade comme on le pensait auparavant. Cette observation semble plutôt indiquée que ce clade aurait probablement existé au moins au Moyen-Orient, avant que le contact entre les amérindiens et les européens s’est établi. Ce résultat supporte donc une origine asiatique pour ce clade, suivi par son introduction dans le Nouveau Monde par les premiers hommes ayant atteint le continent américain par le détroit de Béring il y a des milliers d’années, où les poux sont restés in-situ jusqu'au deuxième contact avec les colons européens au 16ème siècle.
Dans le deuxième travail, nous avons analysé des poux de tête récoltés sur des amérindiens Wayampi de village trois-saut d’Amazonie. L’analyse de l’ADN mitochondrial ciblant trois gènes a révélé l’appartenance probable de ces poux à un nouveau clade mitochondrial que nous avons nommé clade « F ».
45 / 285
Article 2 :
High Ancient Genetic Diversity of Human Lice, Pediculus humanus, from Israel Reveals New Insights into the Origin of Clade B Lice
Publié dans PloS One. 2016;11: e0164659
46 / 285 RESEARCH ARTICLE High Ancient Genetic Diversity of Human Lice, Pediculus humanus, from Israel Reveals New Insights into the Origin of Clade B Lice
Nadia Amanzougaghene1, Kosta Y. Mumcuoglu2, Florence Fenollar1,3, Shir Alfi2, Gonca Yesilyurt2, Didier Raoult1,3, Oleg Mediannikov1,3*
1 Unite´ de Recherche sur les Maladies Infectieuses Tropicales Emergentes (URMITE) UM63, CNRS 7278, IRD 198, INSERM 1095, Aix-Marseille Universite´, Marseille cedex 05, France, 2 Parasitology Unit, Department of Microbiology and Molecular Genetics, The Kuvin Center for the Study of Infectious and Tropical Diseases, Hadassah Medical School, The Hebrew University, Jerusalem, Israel, 3 Campus International UCAD-IRD, Dakar, Senegal
* a11111 [email protected]
Abstract
The human head louse, Pediculus humanus capitis, is subdivided into several significantly divergent mitochondrial haplogroups, each with particular geographical distributions. His- OPEN ACCESS torically, they are among the oldest human parasites, representing an excellent marker for Citation: Amanzougaghene N, Mumcuoglu KY, tracking older events in human evolutionary history. In this study, ancient DNA analysis Fenollar F, Alfi S, Yesilyurt G, Raoult D, et al. (2016) using real-time polymerase chain reaction (qPCR), combined with conventional PCR, was High Ancient Genetic Diversity of Human Lice, applied to the remains of twenty-four ancient head lice and their eggs from the Roman Pediculus humanus, from Israel Reveals New Insights into the Origin of Clade B Lice. PLoS ONE period which were recovered from Israel. The lice and eggs were found in three combs, one 11(10): e0164659. doi:10.1371/journal. of which was recovered from archaeological excavations in the Hatzeva area of the Judean pone.0164659 desert, and two of which found in Moa, in the Arava region, close to the Dead Sea. Results Editor: Feng Gao, Tianjin University, CHINA show that the head lice remains dating approximately to 2,000 years old have a cytb hap-
Received: July 8, 2016 logroup A, which is worldwide in distribution, and haplogroup B, which has thus far only been found in contemporary lice from America, Europe, Australia and, most recently, Accepted: September 28, 2016 Africa. More specifically, this haplogroup B has a B36 haplotype, the most common among Published: October 14, 2016 B haplogroups, and has been present in America for at least 4,000 years. The present find- Copyright: © 2016 Amanzougaghene et al. This is ings confirm that clade B lice existed, at least in the Middle East, prior to contacts between an open access article distributed under the terms Native Americans and Europeans. These results support a Middle Eastern origin for clade of the Creative Commons Attribution License, which permits unrestricted use, distribution, and B followed by its introduction into the New World with the early peoples. Lastly, the pres- reproduction in any medium, provided the original ence of Acinetobacter baumannii DNA was demonstrated by qPCR and sequencing in four author and source are credited. head lice remains belonging to clade A. Data Availability Statement: All sequences of cytb haplotypes are available in GenBank under accession numbers KX232678-KX232679 and KX249763-KX249775.
Funding: The authors received no specific funding Introduction for this work. The human louse, Pediculus humanus, is an obligatory haematophagous parasite that thrived Competing Interests: The authors have declared exclusively on human blood for at least 5–7 million years ago [1, 2]. This species includes two that no competing interests exist. ecotypes: the head louse (Pediculus humanus capitis De Geer) that lives and multiplies on the
PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 1 / 14 47 / 285 Pediculus humanus remains from Roman-Era
scalp, and the body louse (Pediculus humanus humanus Linnaeus), that lives and oviposits on clothes [3, 4, 5]. Until recently, only the body louse was assumed to act as a vector for at least three serious human diseases, namely epidemic typhus, trench fever and louse-borne relapsing fever caused by Rickettsia prowazekii, Bartonella quintana, and Borrelia recurrentis, respectively [6]. Body lice have also been shown to be able to host and possibly transmit the agent of plague, Yersinia pestis [5, 7]. Though head lice have been found in nature to carry the DNA of Bartonella quin- tana, Borrelia recurrentis, Acinetobacter baumannii and Yersinia pestis [5, 8, 9, 10, 11, 12, 13], and experimental infections have shown that these lice can also act as a vector of louse-borne diseases [14, 15], their epidemiological significance is still debated. Mitochondrial DNA (mtDNA) has been widely used to study the genetic diversity of human lice, revealing the presence of several deeply divergent mtDNA clades or haplogroups named A, B and C [1, 2, 16, 17, 18]. Haplogroup A is the most common. It can be found throughout the world and includes both head and body lice [1, 2, 17, 18]. Clade B comprises only head lice. It is confined to the New Word, Europe and Australia and was recently reported from North and South Africa [2, 18, 19, 20]. Clade C, which only includes head lice, has been found in Ethiopia, Nepal and Thailand [13, 16]. Most recently, two additional novel clades were described in 2015 by Drali et al. and Ashfaq et al. [5, 20] besides the three classical recog- nized clades. The first novel clade is the clade D described by Drali et al. [5] and is referred as clade E in Ashfaq et al. [20]. This clade (clade D sensu Drali et al.) is the sister group of clade A and consist on lice from Ethiopia and Democratic Republic of the Congo and comprising both head and body lice [5]. The second novel clade is described only by Ashfaq et al. [20]. This clade is the sister group of clade C and consist on lice from Senegal and Mali, referred here as clade “E”. This clade comprises only head lice. Pediculus humanus is probably one of the oldest and most intimate human parasites [21, 22] and is known to have a long-term co-evolutionary association with humans over millions of years [1]. As such it represents an excellent marker for tracking older events in human evo- lutionary history [4, 18]. Louse infestations are mentioned in the Bible as the third plague [21]. Lice have also been found in a variety of archaeological contexts around the word [21, 22, 23, 24]. These reports all indicate the long-time presence of lice throughout the world before the globalization initiated during Columbus’s era, as the result of early human migrants out of Africa [4]. For example, in the Old World, the earliest record of head lice goes back to the Neolithic age, roughly 9,000 years ago, obtained from an individual who lived in the Nahal Hemar cave in Israel [21]. In the New World, the oldest such record of head lice comes from an archaic human skeleton in Bra- zil, dated at more than 10,000 years old [22]. However, few molecular data on ancient lice are available. Indeed, ancient DNA offers a direct means to assess the past genetic structure and diversity of human lice, which can provide relevant information relating to the antiquity of migration patterns of humans, their lice and, therefore, the flow of louse-borne pathogens [25]. Thus, based on molecular analyses of head lice from Peruvian mummies, D. Raoult et al. showed that the worldwide clade A had a pre-Columbian presence in the American continent and likely had links to the Old World [17]. In 2013, A. Boutellis et al. confirmed this result and demonstrated that Clade B was also present in America for more than 4,000 years, prior to con- tact with Europe, suggesting an American origin for this haplogroup, followed by its dispersal to the Old World from America beginning in the 16th century [26]. Despite this finding, the precise source of this haplogroup prior to globalization remains unclear. However, other stud- ies have suggested that could originate from Asia, which is reported to have populated the Americas [1, 18].
PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 2 / 14 48 / 285 Pediculus humanus remains from Roman-Era
The analysis of ancient head louse eggs recovered from Israel dating from the Chalcolithic and early Islamic period, showed that these eggs may have belonged to people originating from West Africa, based on identification of the louse mitochondrial sub-clade C specific to that region [25]. In the present study, ancient DNA analysis of head lice remains dating from Roman period recovered from Israel was undertaken in order to: (1) identify their mitochondrial phylotypes, (2) reveal the phylogenetic relationship between ancient and contemporary human lice, and (3) look for louse-borne pathogens in these remains.
Materials and Methods Ancient Head Lice Remains Head lice and their eggs were isolated from three louse combs from the Roman period (1st cen- tury AD to 6th century BC). Comb A and Comb B were recovered from archaeological excava- tions in Moa, in the Arava region, close to the Dead Sea, while Comb C was found in the Hatzeva area of the Judean desert (between Jericho and Dead Sea). In Moa, the excavations were conducted between 1981 and 1985 (permit no. A-1016/1981). The site appears to be a way station on the Spice Route connecting Petra to Gaza, where the remains of a caravanserai, a fortress and a temple were found. The excavation site in Hatzeva was a fortified road station from the Nabatean period in an agricultural settlement from the Early Arabic, late Roman period. The three combs were two-sided (Fig 1). From comb A, six lice parts and four eggs were isolated, from comb B, one entire specimen of male and nymph, respectively, eight lice parts and three eggs while from comb C only one entire nymph was isolated. All of the lice remains were preserved dry under sterile conditions. Morphological examination revealed that
Fig 1. Recovery of ancient human head lice from a two-sided louse comb belonging to the Roman period (A) recovered from the Judean desert and Arava regions of Israel. In the lower part, entire specimens (B and C), the head and thorax of a head louse (D) and a damaged non-operculated egg (E) can be seen. doi:10.1371/journal.pone.0164659.g001
PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 3 / 14 49 / 285 Pediculus humanus remains from Roman-Era
most of the eggs were embryonated. All examined combs are deposited in the Antiquities Authority in Jerusalem, Israel. Permission has been obtained from the Antiquities Authority to examine the combs and publish the results. All necessary permits were obtained for the described study, which complied with all relevant regulations.
Ancient DNA Analysis DNA extraction. Lice and eggs were rinsed twice in distilled water for 15 minutes and then crushed individually in sterile Eppendorf tubes. Two extractions blank were systematically co-extracted with the ancient lice samples during each extraction session. No more than five ancient samples were co-extracted at the same time. Finally, the total-DNA was extracted from seventeen head lice parts and seven nit/egg-parts using a phenol/chloroform/isoamyl protocol [27]. The extracted genomic DNA concentrations varied between 1.2 to 1.7 ng/ml for seven nit/egg-parts and between 1.4 to 2.4 ng/ml for seventeen head lice parts. Identification of lice DNA and determination of lice clade. In order to decrease the pos- sibility of contamination and false-positives, two sensitive sets of primers and probes for real- time quantitative polymerase chain reaction (qPCRs) were specifically designed for this study. Both qPCRs were designed in order to amplify all known clades of P. humanus, targeted 88-bp and 100-bp fragments of the cytochrome b gene (cytb) and the 12S ribosomal RNA (12S RNA), respectively. The design was performed with Primer3 software, version 4.0 (http://frodo.wi. mit.edu/primer3/), according to procedures described elsewhere [28]. The oligonucleotide sequences of the primers and probes were as follows: CytbF1 (5'- AGTGCTATTCCTRTTRTT GG-3'), CytbR1 (5'-AAYARYCGCTCTAAAGT AGG-3') and TaqMan probe (FAM-TGAGGA GGGTTTTCAGT- MGB) for the cytb gene, 12SF1 (5'- ATCTTACCTTTTAACTTTTGCT-3'), 12SR1 (5'- GCGTCTTGACTTGTACRTTA-3') and TaqMan probe (FAM- CTGGCACGTCG CTGTACTAA—MGB) for the 12S ARN gene. PCR amplification was performed using a CFX96 Thermal Cycler (Bio-Rad Laboratories, Foster City, CA, USA) in a 20 μl reaction volume containing 5 μL of the DNA template, 10 μL of Eurogentec™ Probe PCR Master Mix (Eurogentec, Liège, Belgium), 0.5 μM of each primer and 0.5 μM of the FAM-labeled probe. The thermal cycling conditions included one incubation step at 50°C for two minutes and an initial denaturation step at 95°C for three minutes, fol- lowed by 40 cycles of denaturation at 95°C for 15 seconds and annealing extension at 60°C for 30 seconds. At first, all the samples were screened in both qPCRs. Thereafter all those which were posi- tive by at least one qPCR were subjected to conventional PCR targeting the 272 bps portion of cytb gene as described by Boutellis et al.[19]. All the products of the PCR amplification were checked on gel electrophoresis and then purified using NucleoFast 96 PCR plates (Macherey-Nagel EURL, Hoerdt, France) according to the manufacturer’s instructions. All products of both qPCRs and conventional PCR were sequenced with original primers on an ABI automated sequencer (Applied Biosystems, USA) with the BigDye Terminator v1.1 cycle (Applied Biosystems, Foster City, CA). The electrophe- rograms obtained were assembled and corrected using ChromasPro software (Technelysium Pty, Queensland, Australia). Detection of pathogens. To test for the presence of pathogens, qPCRs were performed using previously reported primers and probes targeting the 16S rRNA gene of Borrelia [29], the ITS intergenic spacer of Bartonella [8], the rpoB gene of Acinetobacter [9], the ompB gene of Rickettsia prowazekii and the pla gene of Yersinia pestis [27]. All qPCRs were performed using a CFX96 TMREAL-Time System C1000 Thermal Cycler (Bio-Rad Laboratories) and the Euro- gentec Master Mix Probe PCR kit (Eurogentec).
PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 4 / 14 50 / 285 Pediculus humanus remains from Roman-Era
Prevention of DNA contamination. In order to ensure that no contamination by modern DNA would interfere with the results, all pre-PCR and post-PCR procedures were performed respectively in a separate, clean room, free of louse DNA under a hood with air-capture, using autoclaved and UV treated material. Extractions and PCR amplification blanks were used as negative controls, in order to detect possible contamination by external DNA. No positive con- trol was included in any experimental steps in order to minimize potential contamination. No amplifications were detected among the negative controls throughout the study.
Data Analysis MEGA 6.06 was used for phylogenetic analyses and tree reconstruction with 500 bootstrap rep- licates using the maximum likelihood method under Kimura’s 2-parameter with complete deletion [30] For comparison, the ancient DNA sequences obtained in this study were combined with the 30 cytb haplotypes reported by Drali et al.[31]. This dataset was complemented with newly available sequences in GenBank, after being assigned to haplotypes using DnaSP v5.10 [32]. Ancient lice sequences from 10,000 and 4,000 year-old Peruvian and Chilean mummies, respectively, were also included [17, 26]. Finally, a dataset that consisted of 49 haplotypes was created in which two are shared between contemporary and ancient lice. These haplotypes span 40 geographic locations (countries) on five continents (S1 Table). The novel haplotypes identified were deposited in GenBank under the following accession numbers: KX249763-KX249775. In order to investigate the possible relationships among haplotypes, the median-joining (MJ) network using the Bandelt method was constructed using the NETWORK4.6 program (www.fluxus-engineering.com/sharenet.htm)[33]. For A. baumannii, the nucleotide sequences obtained in this study were aligned with the ref- erence sequences available in public databases (GenBank) and a phylogenetic tree was also con- structed using the maximum likelihood method under Kimura’s 2-parameter model implemented in MEGA 6.0 6, with 500 bootstrap replicates [30].
Results In this study, we isolated DNA from twenty-four head lice parts and their eggs, P. h. capitis, from Roman-era remains retrieved in Israel. A 110-bp DNA fragment of the 12S RNA gene (qPCR) and 85-bp (qPCR) and 270-bp (conventional PCR) DNA fragments of cytb gene were targeted.
Detection of Lice DNA in Ancient Lice Samples The qPCR targeting an 85-bp fragment of cytb gene was positive for all the 24 ancient DNA lice tested (32 Determination of Lice Clade To determine lice clade, all samples which were positive for at least one qPCR (24/24) were subjected to conventional PCR targeting a 272-bp fragment of the cytb gene coupled with sequencing (S2 Fig). To insure the reproducibility of the results, we also sequenced the prod- ucts of both qPCRs, since the two shorts fragments amplified by both qPCRs can discriminate between all known clades of lice. Thus, at least one sequence per sample was successfully recov- ered from all the samples (24/24). PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 5 / 14 51 / 285 Pediculus humanus remains from Roman-Era In comparison with previously well-defined lice mtDNA haplogroups A, C [16], B [1], D (sensu Drali et al.) [5] and E [20], all ten samples recovered from comb A belonged to mito- chondrial clade B (41.6%). All lice samples from combs B (thirteen samples) and C (one sam- ple) belonged to clade A (58.3%) (Table 1). Phylogenetic Analysis and Haplotype Assignment For the phylogenetic analysis we used the maximum likelihood method (ML) (Fig 2) and a median-joining (MJ) network (Fig 3) analysis. Only the 272-bp sequences of the cytb gene (20/24) were analyzed and combined with all known modern and ancient haplotypes generated in our dataset. Table 1. Summary of ancient samples, DNA analyses and haplotypes assignment. Louse number (Lab Part of lice PCR results Haplogroup Haplotype identity on the basis of partial code) amplified 12S Cytb Cytb identity cytb 270-bp (110-bp) (80-bp) (270-bp) Comb A Romanic-HL1 thorax /abdomen + + + B Hap_B36 Romanic-HL2 thorax /abdomen + + + B Hap_B36 Romanic-HL3 Thorax + + + B Hap_B36 Romanic-HL4 Abdomen + + + B Hap_B36 Romanic-HL5 Abdomen + + + B Hap_B36 Romanic-HL6 Abdomen + + + B Hap_B36 Romanic-HN7 non operculated + + + B Hap_B36 egg Romanic-HN8 non operculated + + + B Hap_B36 egg Romanic-HN9 operculated egg NA + NA B* — Romanic-HN10 operculated egg + + NA B* — Comb B Romanic-HL11 entire male + + + A Hap_A5 Romanic-HL12 entire nymph + + + A Hap_A55 Romanic-HL13 leg/thorax/ + + + A Hap_A55 abdomen Romanic-HL14 thorax /abdomen + + + A Hap_A5 Romanic-HL15 thorax /abdomen + + + A Hap_A5 Romanic-HL16 thorax /abdomen + + + A Hap_A5 Romanic-HL17 Thorax + + + A Hap_A55 Romanic-HL18 Thorax + + + A Hap_A5 Romanic-HL19 abdomen + + + A Hap_A55 Romanic-HL20 abdomen + + + A Hap_A5 Romanic-HN21 operculated egg NA + NA A* — Romanic-HN22 non-operculated + + + A Hap_A5 egg Romanic-HN23 non-operculated + + NA A* — egg Comb C Romanic-HL24 entire nymph + + + A Hap_A56 Total 24 22/24 24/24 20/24 14(A)+10(B)/24 7(A5)+4(A55)+ 1(A56)+8(B36)/24 NA: not amplified *- clades identified on the base of cytb qPCR product sequencing doi:10.1371/journal.pone.0164659.t001 PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 6 / 14 52 / 285 Pediculus humanus remains from Roman-Era Fig 2. Cytb haplotype networks of contemporary and ancient human body and head lice. Each circle area indicates a unique haplotype and variations in circle size are proportional to haplotype frequencies. Pie colors and sizes in circles represent the continents and the number of their sequence for a haplotype. The length of the links between nodes is proportional to mutational differences. doi:10.1371/journal.pone.0164659.g002 Within clade A (12/20), seven sequences (all from comb B) of these ancient head lice belonged to the widespread modern haplotype A5. This haplotype is the most prevalent worldwide (75% of locations and 45% of the 985 analyzed human lice) and is present in all continents. It occupies a central position in haplogroup A, as shown in the MJ network, and is also found in pre- Columbian Peruvian and Chilean mummies from the New World. The five remaining clade A sequences, four from comb B and one from comb C, yielded two novel haplotypes which were found to be unique, provisionally named here as A55 and A56, respectively. A thorough inspec- tion of the haplotype network revealed that the A55 haplotype deviated from haplotype A5 by one mutation step while the A56 haplotype derived on haplotype A5 by two mutations steps. Nevertheless, the mutations that we found in both haplotypes A55 and A56 are synonymous, i.e. resulting in no amino acid changes, compared to the A5 haplotype. The sequences of these two novel haplotypes were deposited in GenBank under accession number: KX232678-KX232679. Within the clade B ancient head lice, all eight sequences (all from comb A) belonged to the modern haplotype B36, which is the most common within the B-haplogroup (74% of the 184 ana- lyzed haplogroup B human lice). This haplotype occupied a central position in the MJ network, and is found in Africa, Europe and America, as well as in pre-Columbian Chilean mummies. Detection of Pathogens The qPCR targeting the rpoB gene of Acinetobacter was positive for four samples (4/24) of ancient lice DNA tested (34 PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 7 / 14 53 / 285 Pediculus humanus remains from Roman-Era Fig 3. Maximum-likelihood (ML) phylogram of contemporary and ancient haplotypes of Pediculus humanus based on the partial 272-bp cytb gene with Pediculus schaeffi (KC241883) and Pthirus pubis (EU219990) as outgroups. doi:10.1371/journal.pone.0164659.g003 directly sequenced. The four obtained sequences of the partial rpoB gene (182-bp) were 100% identical to one another and were identified as A. baumannii on the basis of a BLAST search. These sequences had 180 of 182 base positions in common (98.9% identity) with a reference strain of A. baumannii (GenBank accession number CP012952) and 181 of 182 base positions in common (99.4% similarity) with A. baumannii isolated from modern human head lice PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 8 / 14 54 / 285 Pediculus humanus remains from Roman-Era Fig 4. Acinetobacter baumannii from ancient head lice belonging to the Roman period. a, Maximum-likelihood (ML) phylogenetic tree relationship based on 182-bp fragment rpoB gene of A. baumannii detected in ancient head lice was compared with the reference sequences strain, while Pseudomonas was used as an out group. Bootstrap values are indicated at the nodes. Bold indicates the taxonomic position of A. baumannii identified in this study. b, 182-bp of A. baumannii rpoB gene fragment sequenced from the four ancient head lice, exhibiting one mutation not present in the homologous gene sequence from its closest relative, the modern A. baumannii sequence in GenBank”. doi:10.1371/journal.pone.0164659.g004 collected from elementary school children in Thailand (GenBank accession number KP161047). However, these ancient sequences have one mutation (position 81) that has not previously been described in modern A. baumannii (Fig 4B). The phylogenetic tree of this A. baumannii is shown in Fig 4A. All four A. baumannii positive ancient head lice belonged to clade A. Borrelia spp., Bartonella spp., Rickettsia prowazekii and Y. pestis DNA could not be identi- fied in any of the 24 ancient head lice specimens tested. PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 9 / 14 55 / 285 Pediculus humanus remains from Roman-Era Discussion A paramount concern in studying ancient DNA is the prevention of its contamination by mod- ern DNA [34]. Several elements support the authenticity of the results presented herein: qPCRs were designed specifically for this study, each experimental step was performed in sepa- rate rooms, free of lice and their DNA and no positive controls were used in the PCR examina- tions as recommended elsewhere [34]. Some of the sequences obtained were unique and presented mutations which have never previously been reported, lending greater confidence to the authenticity of the present results [34]. The mtDNA analysis in this study revealed that the head lice remains from approximately 2,000 years ago have cytb haplogroups A and B. To the best of our knowledge, the present study is the first to reveal the presence of a mitochondrial haplogroup B in a Middle Eastern region. It has thus far only been found in contemporary lice from America, Europe, Australia and, more recently, Africa (Algeria and South Africa) [2, 18, 19, 20]. Specifically, this hap- logroup B has a B36 haplotype, the most common among B haplogroups [31]. Interestingly, this haplotype was also present for at least 4,000 years in South America before the arrival of European settlers [26]. Accordingly, the present findings show that clade B lice existed in the Mediterranean region long before the discovery of America, meaning that clade B lice did not originate from America as was previously thought but probably existed, at least in the Middle East, prior to contact between Native Americans and Europeans. These results are consistent with the hypothesis suggesting that this clade could originate from Asia, which is reported to have populated the Americas [1, 18]. The remaining samples had haplogroup A and yielded three haplotypes, in which two hap- lotypes (provisionally called A55 and A56 in this paper) were unique to the ancient head lice examined in this study, while haplotype A5, which is common worldwide, including in ancient head lice from Chilean and Peruvian mummies of the New World [17, 26, 31]. Prior research has suggested that the known lice clades evolved on different lineages of Homo, similarly to those known to date from 2.3 to 0.03 million years ago (MYA) [1, 20] and, accordingly, their geographic distribution can provide information regarding the evolutionary history of the lice as well as their human hosts [2]. Clade A lice most likely emerged in Africa and evolved on the host lineage that led to anatomically modern humans (Homo sapiens), showing signs of a recent demographic expansion out of Africa about 100,000 years ago, first to Eurasia and subsequently to Europe, Asia and the New World [1, 18]. Haplogroup B diverged from haplogroup A between 0.7 and 1.2 million years ago and may have evolved on archaic hominids, such as the Homo neanderthalensis, who expanded in Europe and Asia, and only became associated with modern humans during the period of overlap as the result of a recent host switch [1, 3, 4]. Thus, considering the present-day geographic distribution of the two haplotypes A5 and B36 and evidence of their long-time presence in the New World, along with our data from the Middle East, the most likely theory is that these two haplotypes were carried by early humans and migrated with them throughout the world before the globalization initiated during the time of Columbus. It seems that the B36 haplotype was originally present in archaic populations of the Middle East, and because this region was a passageway for Homo sapiens between Africa and the rest of the world [35], this haplotype could have switched to anatomically modern humans when they arrived, and migrated with them along with lice of haplotype A5 throughout the world including to America, where lice remained in situ for thou- sands of years until the second contact with European colonists in the 16th century [2]. Aside from their role as biological markers used to infer human evolutionary history, lice remains can provide information relating to past human sanitary conditions and diseases, PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 10 / 14 56 / 285 Pediculus humanus remains from Roman-Era because these lice are vectors of bacterial pathogens. Yet, genetic traces from pathogens can be identified in archaeological remains and ancient DNA has successfully been used to identify B. quintana in 4,000-year-old teeth [36] and was reported to occur in lice at the end of World War I [37]. Raoult et al.[34] showed that Napoleon’s soldiers in Vilnius were exposed to body lice containing B. quintana and that soldiers had evidence of infection with either R. prowazekii or B. quintana, concluding that louse-borne infectious diseases affected nearly one-third of Napoleon’s soldiers buried in Vilnius, and indicating that these diseases might have been a major factor in the French retreat from Russia. This study reveals for the first time the presence of A. baumannii DNA in ancient human head lice remains belonging to clade A. Specifically, the rpoB sequence that was found has not been reported previously. In recent years, A. baumannii was first isolated from the body lice of homeless people in Marseille (France) as well as from diverse countries worldwide [38]. A. baumannii DNA was also found in head lice collected from elementary school children in Paris belonging to clade A [9] and detected in body and head lice collected from healthy individuals from Ethiopia [10]. In 2015, S. Sunantaraporn et al. showed that A. baumannii could be detected in clade A and C head lice collected from elementary school children in Thailand [13]. Although A. baumannii was reported to occur in temporary head lice in several occasions, the clinical significance of this finding is unknown [9, 39]. More recently, A. baumannii was shown to cause nosocomial infections and severe community-acquired infections such as pneumonia, bacteremia, endocarditis, and meningitis, due to its increasing resistance to a wide range of antibacterial agents [40, 41]. Conclusion The present work confirms that clade B lice existed, at least in the Middle East, prior to contacts between Native Americans and Europeans. Our results cannot support the previous hypothesis that clade B has an American origin and was imported from America to the Old World after its discovery by Christopher Columbus. We also identified the presence of nosocomial pathogen, A. baumannii in Roman-era head lice remains belonging to clade A. Further study of the lice remains would be necessary to shed more light on the patterns of human migration worldwide, their lice and the flow of louse-borne pathogens at different times in history. Supporting Information S1 Fig. Real-time PCR amplification cuves for ancient DNA, Pediculus humanus, using cytb and 12S gens with negative controls (NC). 1, 2 and 3 showed qPCR amplification tar- geted a 88-bp DNA fragment of cytb gene (24/24 positive with Ct varied between 32 to 38); 4 showed qPCR amplification targeted a 100-bp DNA fragment of 12S gene (22/24 positive with Ct varied between 32 to 38). (PPTX) S2 Fig. Agarose gel stained with syber safe showing the 277-bp amplification for Pediculus humanus cytb gene fragment obtained from ancient DNA with negative controls (T-). (PPTX) S1 Table. Geographic occurrences and frequencies of cytb haplotypes of human head and body lice. Haplotypes highlighted in blue are the newly identified haplotypes from sequences available in GenBank. (XLSX) PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 11 / 14 57 / 285 Pediculus humanus remains from Roman-Era Acknowledgments We would like to thank Dr. Orit Shamir and Dr. Naama Sukenik from the Israel Antiquities Authority, as well as Professor Simcha Lev-Yadun from Haifa University, for their help in iden- tifying and allowing us to examine the ancient louse combs for louse remains. We also thank IHU Méditerranée Infection for financially supporting the study. Author Contributions Conceptualization: NA KYM OM DR FF. Funding acquisition: DR. Investigation: NA KYM OM DR FF SA GY. Methodology:KYM OM DR FF. Project administration: OM DR FF. Resources: OM DR FF. Supervision: OM DR FF. Validation: NA KYM OM DR FF. Writing – original draft: NA OM. Writing – review & editing: NA KYM OM DR FF SA GY. References 1. Reed DL, Smith VS, Hammond SL, Rogers AR, Clayton DH. Genetic analysis of lice supports direct contact between modern and archaic humans. PLoS Biol 2004; 2:e340. doi: 10.1371/journal.pbio. 0020340 PMID: 15502871 2. Light JE, Allen JM, Long LM, Carter TE, Barrow L, Suren G, et al. Geographic distributions and origins of human head lice (Pediculus humanus capitis) based on mitochondrial data. J Parasitol 2008; 94:1275–1281. GE-1618[pii]; doi: 10.1645/GE-1618.1 PMID: 18576877 3. Ascunce MS, Toups MA, Kassu G, Fane J, Scholl K, Reed DL. Nuclear genetic diversity in human lice (Pediculus humanus) reveals continental differences and high inbreeding among worldwide popula- tions. PLoS One 2013; 8:e57619. doi: 10.1371/journal.pone.0057619 PMID: 23460886 4. Boutellis A, Abi-Rached L, Raoult D. The origin and distribution of human lice in the world. Infect Genet Evol 2014; 23:209–217. doi: 10.1016/j.meegid.2014.01.017 PMID: 24524985 5. Drali R, Shako JC, Davoust B, Diatta G, Raoult D. A new clade of African body and head lice infected by Bartonella quintana and Yersinia pestis—Democratic Republic of the Congo. Am J Trop Med Hyg 2015; 93: 990–993. doi: 10.4269/ajtmh.14-0686 PMID: 26392158 6. Raoult D, Roux V. The body louse as a vector of reemerging human diseases. Clin Infect Dis 1999; 29:888–911. doi: 10.1086/520454 PMID: 10589908 7. Houhamdi L, Lepidi H, Drancourt M, Raoult D. Experimental model to evaluate the human body louse as a vector of plague. J Infect Dis 2006; 194:1589–1596. doi: 10.1086/508995 PMID: 17083045 8. Angelakis E, Diatta G, Abdissa A, Trape JF, Mediannikov O, Richet H, et al. Altitude-dependent Barto- nella quintana genotype C in head lice, Ethiopia. Emerg Infect Dis 2011; 17:2357–2359. doi: 10.3201/ eid1712.110453 PMID: 22172306 9. Bouvresse S, Socolovschi C, Berdjane Z, Durand R, Izri A, Raoult D, et al. No evidence of Bartonella quintana but detection of Acinetobacter baumannii in head lice from elementary school children in Paris. Comp Immunol Microbiol Infect Dis 2011; 34:475–477. doi: 10.1016/j.cimid.2011.08.007 PMID: 21974965 10. Kempf M, Abdissa A, Diatta G, Trape JF, Angelakis E, Mediannikov O, et al. Detection of Acinetobac- ter baumannii in human head and body lice from Ethiopia and identification of new genotypes. Int J Infect Dis 2012; 16:e680–e683. doi: 10.1016/j.ijid.2012.05.1024 PMID: 22771379 PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 12 / 14 58 / 285 Pediculus humanus remains from Roman-Era 11. Boutellis A, Mediannikov O, Bilcha KD, Ali J, Campelo D, Barker SC, et al. Borrelia recurrentis in head lice, Ethiopia. Emerg Infect Dis 2013; 19:796–798. doi: 10.3201/eid1905.121480 PMID: 23648147 12. Sangare AK, Boutellis A, Drali R, Socolovschi C, Barker SC, Diatta G, et al. Detection of Bartonella quintana in African body and head lice. Am J Trop Med Hyg 2014; 91:294–301. doi: 10.4269/ajtmh.13- 0707 PMID: 24935950 13. Sunantaraporn S, Sanprasert V, Pengsakul Th, Phumee A, Boonserm, Tawatsin A, et al. Molecular survey of the head louse Pediculus humanus capitis in Thailand and its potential role for transmitting Acinetobacter spp. Parasit Vectors 2015; 8:127. doi: 10.1186/s13071-015-0742-4 PMID: 25889008 14. Goldberger J, Anderson JF. The transmission of Typhus fever, with especial reference to transmission by the head louse (Pediculus capitis). Public Health Reports (1896–1970) 1912; 27:297–307. doi: 10. 2307/4567527 15. Murray ES, Torrey SB. Virulence of Rickettsia prowazekii for head lice. Ann N Y Acad Sci 1975; 266: 25–34. doi: 10.1111/j.1749-6632.1975.tb35086.x PMID: 829471 16. Kittler R, Kayser M, Stoneking M. Molecular evolution of Pediculus humanus and the origin of clothing. Curr Biol 2003; 13:1414–1417. PMID: 12932325 17. Raoult D, Reed DL, Dittmar K, Kirchman JJ, Rolain JM, et al. Molecular identification of lice from pre- Columbian mummies. J Infect Dis 2008; 197:535–543. doi: 10.1086/526520 PMID: 18254682 18. Ascunce MS, Fane J, Kassu G, Toloza AC, Picollo MI, Gonzalez-Oliver A, et al. Mitochondrial diversity in human head louse populations across the Americas. Am J Physiol Anthropol 2013; 152:118–129. 19. Boutellis A, Bitam I, Fekir K, Mana N, Raoult D. Evidence that clade A and clade B head lice live in sym- patry and recombine in Algeria. Med Vet Entomol 2015; 29: 94–98. doi: 10.1111/mve.12058 PMID: 25346378 20. Ashfaq M, Prosser S, Nasir S, Masood M, Ratnasingham S, Hebert PD. High diversity and rapid diver- sification in the head louse, Pediculus humanus (Pediculidae: Phthiraptera). Nature. Scientific Reports 2015; 5: 14188. doi: 10.1038/srep14188 21. Mumcuoglu K. Pediculus and Pthirus. In: Paleomicrobiology–Past Human Infections. Raoult D & Dran- court M (eds). Springer, Berlin 2008; pp. 215–222. 22. Araujo A, Ferreira LF, Guidon N, Maues Da Serra FN, Reinhard KJ, et al. Ten thousand years of head lice infection. Parasitol Today 2000; 16:269. S0169–4758(00)01694-X[pii]. PMID: 10858638 23. Rivera MA, Mumcuoglu K, Mathney RT, Matheny DG. Head lice eggs, Anthropophthirus capitis, from mummies of the Chinchorro tradition, Camarones 15-D, Northern Chile. Chungara, Revistade Antropo- logia Chilena 2008; 40:31–39. 24. Arriaza B, Orellana NC, Barbosa HS, Menna-Barreto RF, Araujo A, Standen V. Severe head lice infes- tation in an Andean mummy of Arica, Chile. J Parasitol 2012; 98:433–436. doi: 10.1645/GE-2903.1 PMID: 22010860 25. Drali R, Mumcuoglu KY, Yesilyurt G, Raoult D. Studies of ancient lice reveal unsuspected past migra- tions of vectors. Am J Trop Med Hyg 2015; 93:623–625. doi: 10.4269/ajtmh.14-0552 PMID: 26078317 26. Boutellis A, Drali R, Rivera MA, Mumcuoglu KY, Raoult D. Evidence of sympatry of clade A and clade B head lice in a pre-Columbian Chilean mummy from Camarones. PLoS One 2013; 8:e76818. doi: 10. 1371/journal.pone.0076818 PMID: 24204678 27. Nguyen-Hieu T, Aboudharam G, Signoli M, Rigeade C, Drancourt M, Raoult D. Evidence of a louse- borne outbreak involving typhus in Douai, 1710–1712 during the war of Spanish succession. PLoS One 2010; 5:e15405. doi: 10.1371/journal.pone.0015405 PMID: 21060879 28. Thornton B, Basu C. Real-time PCR (qPCR) primer design using free online software. Biochem Mol Biol Educ 2011; 39:145–154. doi: 10.1002/bmb.20461 PMID: 21445907 29. Parola P, Diatta G, Socolovschi C, Mediannikov O, Tall A, Bassene H, et al. Tick-borne relapsing fever borreliosis, rural Senegal. Emerg Infect Dis 2011; 17:883–885. doi: 10.3201/eid1705.100573 PMID: 21529402 30. Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: molecular evolutionary genetics anal- ysis version 6.0. Mol Biol Evol 2013; 30:2725–2729. doi: 10.1093/molbev/mst197 PMID: 24132122 31. Drali R, Abi-Rached L, Boutellis A, Djossou F, Barker SC, Raoult D. Host switching of human lice to new world monkeys in South America. Infect Genet Evol 2016; 39:225–231. doi: 10.1016/j.meegid. 2016.02.008 PMID: 26867815 32. Librado P, Rozas J. DnaSP v5: a software for comprehensive analysis of DNA polymorphism data. Bioinformatics 2009; 25:1451–1452. doi: 10.1093/bioinformatics/btp187 PMID: 19346325 33. Bandelt HJ, Forster P, Rohl A. Median-joining networks for inferring intraspecific phylogenies. Mol Biol Evol 1999; 16:37–48. PMID: 10331250 PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 13 / 14 59 / 285 Pediculus humanus remains from Roman-Era 34. Raoult D, Dutour O, Houhamdi L, Jankauskas R, Fournier PE, Ardagna Y, et al. Evidence for louse- transmitted diseases in soldiers of Napoleon’s Grand Army in Vilnius. J Infect Dis 2006; 193:112–120. doi: 10.1086/498534 PMID: 16323139 35. Grugni V, Battaglia V, Hooshiar Kashani B, Parolo S, Al-Zahery N, et al. Ancient migratory events in the Middle East: New clues from the Y-chromosome variation of modern Iranians. PLoS One 2012; 7: e41252. doi: 10.1371/journal.pone.0041252 PMID: 22815981 36. Drancourt M, Tran-Hung L, Courtin J, Lumley H, Raoult D. Bartonella quintana in a 4000-year-old human tooth. J Infect Dis 2005; 191:607–611. doi: 10.1086/427041 PMID: 15655785 37. Maurin M, Raoult D. Bartonella (Rochalimaea) quintana infections. Clin Microbiol Rev 1996; 9:273– 292. PMID: 8809460 38. La Scola B, Raoult D. Acinetobacter baumannii in human body louse. Emerg Infect Dis 2004; 10:1671–3. doi: 10.3201/eid1009.040242 PMID: 15498175 39. Veracx A, Raoult D. Biology and genetics of human head and body lice. Trends Parasitol 2012; 28:563–571. doi: 10.1016/j.pt.2012.09.003 PMID: 23069652 40. Fournier PE, Vallenet D, Barbe V, Audic S, Ogata H, et al. Comparative genomics of multidrug resis- tance in Acinetobacter baumannii. PLoS Genet 2006; 2:e7. doi: 10.1371/journal.pgen.0020007 PMID: 16415984 41. Peleg AY, Seifert H, Paterson DL. Acinetobacter baumannii: emergence of a successful pathogen. Clin Microbiol Rev 2008; 21:538–582. doi: 10.1128/CMR.00058-07 PMID: 18625687 PLOS ONE | DOI:10.1371/journal.pone.0164659 October 14, 2016 14 / 14 60 / 285 Article 3 : Mitochondrial diversity and phylogeography of Pediculus humanus, with the description of a new Amazonian Clade F En préparation pour soumission à Infection, Genetics and Evolution 61 / 285 Mitochondrial diversity and phylogeography of Pediculus humanus, with the description of a new Amazonian Clade F. Nadia Amanzougaghene1, Florence Fenollar2, Bernard Davoust1, Félix Djossou3, Muhammad Ashfaq4, Idir Bitam2, 5, Didier Raoult1, Oleg Mediannikov1* 1Aix Marseille Univ, IRD, APHM, MEPHI, IHU-Méditerranée Infection, Marseille, France 2Aix Marseille Univ, IRD, APHM, VITROME, IHU-Méditerranée Infection, Marseille, France 3Service de maladies Infectieuses et Tropicales, Centre Hospitalier de Cayenne, 97306 Cayenne Cedex, Guyane Française, France 4 Biodiversity Institute of Ontario, University of Guelph, Guelph, ON, Canada. 5 Ecole Supérieure des Sciences de l'Aliment et des Industries Agro-Alimentaires, Algiers, Algeria. *Corresponding author : Dr. Oleg MEDIANNIKOV Address: MEPHI, IRD, APHM, IHU-Méditerranée Infection, 19-21 Boulevard Jean Moulin, 13385 Marseille Cedex 05 Tel: +33 (0)4 13 73 24 01, Fax: +33 (0)4 13 73 24 02, E-mail: [email protected], Word abstract count: 215 Word text count: 3,036 62 / 285 Abstract P. humanus is an obligate and highly intimate bloodsucking insect of human, it occurs on two ecotypes, head louse and body louse. In this study, we have analyzed genetic diversity in head lice collected from Amazonian individuals by targeting three mitochondrial genes (cytb, cox1 and 12S). By coupling these results with all available mitochondrial data of P. humanus from over the world, six highly divergent and well-supported monophyletic clades were identified. Five clades corresponding to previously recognized mitochondrial clades A, D, B, C and E, while the sixth (clade F) was novel, showing 5.4%, 3.7% and 3.6% divergence from its nearest neighbor clade B, respectively, at cytb, cox1 and 12S. Interestingly, this novel clade F was recovered only in some head lice sequences from Mexico and Argentina, while it was the dominant lineage found in the Amazonian lice and it may represent Native America louse mitochondrial diversity. Furthermore, P. mjobergi, a New World monkey louse, which is through to be transmitted to monkeys from the first humans reached the American continent thousands of years ago, is also belonged to this clade. The discovery of new Amazonian clade F with the recovery of additional haplotypes within each of the five clades demonstrate that the levels of genetic diversity in P. humanus is higher than previously thought. Keywords: Pediculus humanus, phylogeography, genetic diversity, Clade F, Amazonia 63 / 285 Introduction Sucking lice (Phthiraptera: Anoplura) are obligate blood-feeding ectoparasites of placental mammals and have coevolving with their hosts for at least the last 65 million years ago (MYa) and likely much longer [1,2]. Humans are parasitized by two species of sucking lice, the pubic louse (Pthirus pubis), and head/body lice (Pediculus humanus) [3]. The association between the P. humanus and its human host is an ancient one, extending back at least 6 MYa to the last common ancestor of humans and chimpanzees [3]. P. humanus includes two ecotypes, head lice (P. h. capitis) and body lice (P. h. humanus), that are morphologically and biologically almost similar, but ecologically distinct [3,4]. Head lice are confined to the scalp and feed on human blood every 4–6 hours [4]. Body lice live in clothing and feed less frequently but take more large blood meal than head lice [4]. Aside from their role as pests [5], body lice are the main vectors of at last three serious human pathogens; Rickettsia prowazekii (the causative agent of epidemic typhus), Bartonella quintana (trench fever) and Borrelia recurrentis (relapsing fever) [6]. It once believed that only body lice can transmit disease, recently however, several combined evidences from epidemiological and laboratory studies strongly implicated head lice as vector for infectious agents, although its vectorial capacity is lower compared to body lice [7–14]. The genetic diversity of human lice has been widely studied using the mitochondrial genes (mainly cytochrome b [cytb] and cytochrome oxidase subunit 1 [cox1] genes) revealing the presence of five highly divergent mitochondrial clades (A, D, B, C and E) [3,15–18]. In addition to this inter-clade diversity, human lice also present intra-clade diversity, illustrated by several distinct haplotypes for each clade. Body lice belong to clades A and D, while head lice encompass the full genetic diversity of clades [7,19,20]. Clade A is the most common and widely distributed across all continents, whereas the other clades are geographically restricted [15,19,21]. Clade D is restricted to Africa and is found in DR Congo, the Republic 64 / 285 of Congo, Ethiopia and Zimbabwe [7,17]. Clade B is found in America, Europe, Australia, Algeria, South Africa, Saudi Arabia and was recently recovered among head lice remains from Israel, dated approximately to 2,000 years old [15,16,19,21–23]. Clade C has been found in some African and Asian countries including Ethiopia, the Republic of Congo, Nepal, Pakistan and Thailand [7,16,18,21,24]. Finally, clade E seems to be specific to West African lice, including Senegal and Mali, where it is highly prevalent and was recently identified in head lice from Nigerian refugees arriving in Algeria and from migrant communities living in Bobigny, France [8,25] (Louni et al., Parasites and Vectors, accepted). Lice are highly host specific and fast-evolving parasites that have been evolving in tandem with their primate hosts for a thousand of years [19,26]. Previous studies showed that the time since divergence among the lice clades (around 2 MYa) far predated the time of divergence of modern human around 200,000 years ago [19,27]. These results suggested that different lice clades evolved on other archaic hominids, likely those known from 2.3 to 0.03 MYa (such as Homo erectus, H. neanderthalensis and H. denisovan), and only switched to modern humans during the recent period of overlaps [3,16]. Moreover, the presence of highly divergent clades geographically isolated can yield important information regarding the evolutionary history of the lice as well as their human hosts [19,28]. Therefore, more detailed analysis of genetic diversity in P. humanus and current distributions of its major clades will provide a more detailed picture of parasite evolution and will clarify additional events in our evolutionary history. In the present study, we have obtained and analyzed head lice collected from Amazonian individuals of the Wayampi community living in “Trois-Sauts”, a remote and isolated village. These results are integrated with all prior mitochondrial data from over the world to expand perspectives on the number, distributions, and diversification rates of clades of Pediculus lice. 65 / 285 Materials and methods 1. Ethical approval The Amazonian head lice were collected from infested individuals after getting their verbal consent or from their legal representatives in the case of children, because most of the subjects were illiterate. Local authorities approved and were present when the collection of lice was performed. 2. Louse samples The Amazonian head louse specimens were recovered in 2013 from Amerindians of the Wayampi community living in “Trois-Sauts” (2°15'0" N et 52°52'60" W, 122 m), a remote and isolated village on the Oyapock River along the border between French Guyana and Brazil. In total 98 head lice were recovered from 22 individuals. No body lice were found during the examination. Collected lice were then preserved in 70% ethanol before being sent to our laboratory in Marseille (France). All of the samples were photographed with a camera (Olympus DP71, Rungis, France). In addition, a total of 327 louse specimens were also included in this study, corresponding to body and head lice collected from several countries. These were obtained from the private frozen collection of world lice belonging to our laboratory. 3. DNA extraction, PCR amplification and sequencing Genomic DNA was isolated from louse specimens using the DNeasy tissue kit (Qiagen, Courtaboeuf, France) as described previously [8]. Three mitochondrial genes cytb, cox1 and 12S ribosomal RNA were targeted. PCR amplifications were conducted in a Peltier PTC-200 thermal cycler (MJ Research Inc., Watertown, MA, USA) with the Hotstar Taq-polymerase (Qiagen) in accordance with the manufacturer’s instructions. The success of PCR amplification was then verified by electrophoresis of the PCR product on a 1.5% agarose 66 / 285 gels. All primers used for these experiments and PCR conditions are described in Table 1. Purification of PCR products was performed using NucleoFast 96 PCR plates (Macherey- Nagel EURL, Hoerdt, France) as per the manufacturer’s instructions. The amplicons were sequenced using the Big Dye Terminator Cycle Sequencing Kit (Perkin Elmer Applied Biosystems, Foster City, CA) with an ABI automated sequencer (Applied Biosystems). The electropherograms were assembled and edited using ChromasPro (ChromasPro 1.7, Technelysium Pty Ltd., Tewantin, Australia). 4. Sequences analysis. Cytb, Cox1 and 12S sequences obtained in this study were combined with all available sequences of P. humanus from GenBank database to generate a global dataset to examine clade diversity in P. humanus. The DNA sequences obtained from the literature varied in length, so sequences were trimmed to produce a dataset that maximizes the number of sequences incorporated. The sequences between nucleotide positions 433–705 of cytb (272- bps, according to Genbank Accession KC685778), 748–1031 of cox1 (283-bps, according to Genbank Accession KC685838) and 109-666 of 12S (557-bps, according to Genbank Accession KC685877) were determined for all DNA sequences. ClustalW alignments were performed in MEGA6 [29]. Haplotypes were identified using DnaSP v5.10 software [30]. Finally, we created three datasets for which a total of 1500, 842 and 442 sequences were included, respectively, for cytb, cox1 and 12S datasets. 5. Phylogenetic analysis Neighbor-joining (NJ) analysis was performed in MEGA6 using the K2P model with pairwise-deletion and 500 bootstrap replicates. The Maximum-likelihood (ML) analysis was also performed in MEGA6 using the Kimura 2-parameter model for nucleotide sequences under 500 bootstrap replicates. The subtree for each clade of lice was collapsed with the “compress/expand subtree” function. Cytb, cox1 and 12S sequences from P. schaeffi 67 / 285 (AY695999, KC241887, AY696067, KC241883 and KR706169) were employed as outgroups. 6. Genetic diversity and haplotype analysis For each dataset, population genetic indices including number of sequences (n), number of polymorphic sites (S), average number of pairwise nucleotide differences (k), nucleotide diversity (π), number of haplotypes (H) and haplotype diversity (Hd) were calculated using DNASP v5.10 software [30]. Kimura-2-parameter (K2P) pairwise distances among the cytb, Cox1 and 12S haplotypes were calculated using MEGA6 with pairwise deletion of gaps and missing data [29]. Neutrality tests (Fu & Li’s D and Tajima’s D) were calculated with DNASP v5.10 [30]. In order to investigate the possible relationships between the haplotypes, networks haplotypes for each of the three genes were constructed with the median joining method of Bandelt available in NETWORK5.0 (www.fluxus-engineering.com/sharenet.htm) using equal weights for all mutations [31]. Results A total of 98 head lice collected from 22 Amazonian individuals were analyzed, targeting three mitochondrial genes (cytb, cox1 and 12S). We obtained 11 haplotypes of cytb, 8 haplotypes of cox1 and 13 of 12S that were defined by 31, 25 and 41 polymorphic sites, respectively. The generated Amazonian sequences for each gene were then combined with all the publicly available sequences for cytb, cox1 and 12S, as well as the newly generated cytb, cox1 and 12S sequences in this study (327 sequences). The number of haplotypes in each dataset within each clade and their distributions was determined for each gene. The details of the identified haplotypes, their GenBank accession numbers and geographic locations are described in Table S1-S3. 68 / 285 For cytb dataset 1500 sequences were included from which 105 haplotypes, including one haplotype from P. mjobergi, were identified from 45 countries in the five continents. For cox1 dataset 842 sequences were included from which 57 haplotypes, including one haplotype from P. mjobergi, were identified from 27 countries in the five continents. For 12S dataset 442 sequences were included from which 49 haplotypes, including one haplotype from P. mjobergi, were identified from 18 countries in the five continents. Neighbor-joining (NJ) and Maximum-likelihood (ML) analyses, including all haplotypes was performed for each of the three mtDNA genes, consistently recovered six highly divergent and well-supported monophyletic clades (Fig. 2 and Fig. S1). Five clades corresponding to previously recognized mitochondrial clades A, D, B, C and E, while the sixth was novel, named here “clade F”. This novel clade consists mostly on Amazonian head lice (in total 84 of the 98 [85.7%] Amazonian lice sequences belonged to this clade) as well as sequences from the New World monkey louse P. mjobergi, whereas, the remaining 14 of the 98 (14.3%) Amazonian lice sequences were belonged to clade A. More precisely, for the 12S gene the clade F consisted on 9 haplotypes, 8 haplotypes were from the Amazonian lice (named here F19-F26) from which the haplotype F9 was the most prevalent (83.3 % of 84 sequences), while the ninth haplotype was from P. mjobergi. For the cytb gene, the clade F was also consisted on nine haplotypes, 8 haplotypes were from the Amazonian lice (named here F54 and F1-F7) from which the haplotype F54 was the most prevalent (84.3% of 89 sequences were from the Amazonian head lice sequenced in this study, while 15.7% of 89 sequences also from Amazonian lice and were recovered from Genbank), the remaining haplotype was from P. mjobergi. Lastly, for cox1 gene 8 haplotypes were identified, six haplotypes were from Amazonian lice (named here F29-F34), one 69 / 285 haplotype named F18 was consisted on sequences recovered from GenBank from Argentina (10 cox1 sequences) and Mexico (2 sequences), the eighth haplotype was P. mjobergi. The median-joining networks for all cytb, cox1 and 12S haplotypes corroborated the neighbor joining and Maximum-likelihood phylogenetic reconstructions, with all the recovered haplogroups forming separate clusters represented by six connected subnetworks corresponding to clades A, D, B, C, E and “F” (Fig. 3-5). The maximum intra-clades distances at cytb were 1.2%, 1.9%, 1.4%, 1.4%, 1.5% and 1.1% for clades A (n= 34 haplotypes), D (n= 17), B (n= 9), C (n= 13), E (n= 23) and “F” (n= 8), respectively. The maximum intra-clades distances at cox1 were 1.2%, 1.0%, 1.3%, 1.3%, 1.3% and 1.1% for clades A (n= 17), D (n= 7), B (n= 12), C (n= 6), E (n= 7) and “F” (n= 7), respectively. The maximum intra-clades distances at 12S were 0.9%, 1.6%, 0.4%, 1.4%, 0.3% and 0.5% for clades A (n= 15), D (n= 8), B (n= 3), C (n= 10), E (n= 4) and “F” (n= 8), respectively. The nearest neighbors (NN) distances between clades and the nodal supports are presented in Fig. 2A (cytb), Fig. 2B (cox1) and Fig. 2C (12S). The novel clade “F” shows divergence from its NN clade B of 5.4%, 3.7% and 3.6%, respectively, at cytb, cox1 and 12S. Estimates of genetic diversity indices and the results of neutrality tests for cytb, cox1 and 12S are shown in Table 2. The average number of nucleotide diversity (π), pairwise nucleotide differences (k) and haplotype diversity (Hd) varied among the clades of three genes. The highest haplotype diversity was found within clade D in both cytb and 12S genes (Hd= 0.831 and 0.899, respectively, in cytb and 12S), while in cox1 gene, the highest haplotype diversity was found within clade B (Hd = 0.899). Overall, both (k) and (π) were similar in cytb, cox1 and 12S. 70 / 285 Discussion In this study we have analyzed genetic diversity in head lice collected from Amazonian individuals of the Wayampi community living in “Trois-Sauts”, a remote and isolated village. In total three mitochondrial genes were targeted (cytb, cox1 and 12S). By coupling these results with all available mitochondrial data of P. humanus from over the world, the present study has expanded understanding of its levels and patterning of sequence divergence and revealed higher levels of mtDNA diversity in P. humanus corroborating the results reported by others [16]. To our knowledge, this is the most geographically widespread study to evaluate the genetic diversity in human lice based on three mtDNA genes. Previous studies on P. humanus found that maximum distances within clades of 1.4% at cytb and 1.9% at cox1, while NN distances at these genes were 4.6% and 2.3%, respectively [16]. In the present study maximum distances within clades were almost similar (cytb 1.9%, cox1 1.3% and 1.6% 12S) while NN distances were higher (cytb 5.6% and cox1 6.5%). These results reflect the extended geographic coverage and the larger sample sizes that led to the recovery of additional clade and additional haplotypes within each clade. Three phylogenetic methods (NN, ML and MJ) at three mtDNA genes, consistently recovered six highly divergent and well-supported monophyletic clades. Five clades corresponding to previously recognized mitochondrial clades A, D, B, C and E [15,16], while the sixth (clade F) was novel, consists mainly on head lice from Amazonian individuals analyzed in this study and some sequences of head lice from Argentina and Mexico. Interestingly, the monkey louse P. mjobergi (previously classified within clade B by Drali et al.,[20]) which is through to be transmitted to monkeys from the first humans reached the American continent thousands of years ago [20], is also belonged to this clade. Clade “F” shows divergence from its NN clade B of 5.4%, 3.7% and 3.6%, respectively, at cytb, cox1 and 12S. Clade B was first described in contemporary lice from the 71 / 285 America continent, where it was highly prevalent and diversified [3,19,28]. This finding together with its identification in pre-Columbian mummies’ lice have lead researchers at the beginning to infer an American origin for this clade [32]. However, its recent discover among head lice remains from Israel, dated approximately 2,000 years old, has challenged this assumption and strongly supported an Asian origin for this clade, resulted probably from a recent host switch from Neanderthals to modern humans, followed by its introduction into the New World with the early peoples [3,15,19]. Because clade F was found only in head lice from Mexico and Argentina, and was the dominant lineage found in the Amazonian lice, knowing that Amazonia is one of the few places in the world that has not been strongly affected by globalization, this clade may be the descendants of a pre-Columbian population and was derived from clade B brought by the first humans reached the American continent via the Bering straits thousands of years ago, thus representing Native America louse mitochondrial diversity. Previous studies showed that clade A is the most comment and has a global distribution [8,16,28], results supported by its detection in approximately 46% of the analyzed lice from 49 countries of the five continents. Furthermore, the clade A subnetworks at the three analyzed mitochondrial genes (Fig. 3-5) were star-like in structure, combined with its significant negative Tajima's D value (-2.322, -1.866 and -2.055, respectively at cytb, cox1 and 12S; P<0.05) indicate the signature of population expansion for this clade [33], corroborating the results reported by others [3,28]. In addition, Reed et al. estimated that the demographic expansion of this clade was happened about 100,000 years ago, coinciding with the out-of-Africa expansion of Homo sapiens, thus reflecting a codemographic pattern between lice and humans [3,28]. Clade D (referred as clade E in Ashfaq et al. 2015) diverged from clade A between 0.37 and 0.54 MYa [16] and it is restricted to Central Africa including DR Congo and the Republic of Congo, where it was detected mostly among indigenous 72 / 285 pygmy populations [7,17]. This clade was also reported in lice from Ethiopia [7] and we identified in this study its occurrence, for the first time, in body lice from Zimbabwe. Our sampling did not encounter any new specimens of clade C, so it remains restricted to Africa and Asia [7,19,24,28]. Given its early divergence in the Pediculus tree around 2 MYa, this clade may have evolved on archaic hominids in Asia or Africa such as H. erectus [3,19]. Lastly, clade E (referred as clade D in Ashfaq et al, 2015) diverged from the MRCA of clade C between 0.28 and 0.42 MYa [16]. This clade consists on head lice from West Africa including Senegal and Mali where it was highly prevalent [8] and its recently detection in head lice from Nigerian refugees arriving in Algeria and from migrant communities living in Bobigny (France), is likely a result of a recent migration flows from western African countries [25] (Louni et al., Parasites and Vectors accepted). Conclusion Our study highlights the importance of using mitochondrial genes in analysis of phylogeographic patterns and genetic diversity of P. humanus. Six highly divergent and well- supported monophyletic clades were identified. Five clades corresponding to previously recognized mitochondrial clades A, D, B, C and E, while the sixth “clade F’’ was novel. The novel clade F was mainly found in Amazonia, where it was also shared with the monkey louse P. mjobergi and it may represent Native America louse mitochondrial diversity. The recovery of additional haplotypes within each of the five clades (A, D, C, E and B) and the discovery of new clade F demonstrate that the levels of genetic diversity in P. humanus is higher than previously thought, thus reinforcing the importance of continuing to prospect for and phylogeographically characterize the human lice. 73 / 285 Acknowledgments This study was supported by Méditerranée Infection and the National Research Agency under the program « Investissements d’avenir », reference ANR-10-IAHU-03. 74 / 285 References [1] Durden LA, Musser GG. The sucking lice (Insecta, Anoplura) of the world : a taxonomic checklist with records of mammalian hosts and geographical distributions. Bulletin of the AMNH ; no. 218. Sucking lice and hosts 1994. [2] Light JE, Smith VS, Allen JM, Durden LA, Reed DL. Evolutionary history of mammalian sucking lice (Phthiraptera: Anoplura). BMC Evol Biol 2010;10:292. doi:10.1186/1471-2148-10-292. [3] Reed DL, Smith VS, Hammond SL, Rogers AR, Clayton DH. Genetic Analysis of Lice Supports Direct Contact between Modern and Archaic Humans. PLOS Biol 2004;2:e340. doi:10.1371/journal.pbio.0020340. [4] Veracx A, Raoult D. Biology and genetics of human head and body lice. Trends Parasitol 2012;28:563–71. doi:10.1016/j.pt.2012.09.003. [5] Chosidow O. Scabies and pediculosis. The Lancet 2000;355:819–26. doi:10.1016/S0140- 6736(99)09458-1. [6] Raoult D, Roux V. The body louse as a vector of reemerging human diseases. Clin Infect Dis Off Publ Infect Dis Soc Am 1999;29:888–911. doi:10.1086/520454. [7] Amanzougaghene N, Akiana J, Mongo Ndombe G, Davoust B, Nsana NS, Parra H-J, et al. Head Lice of Pygmies Reveal the Presence of Relapsing Fever Borreliae in the Republic of Congo. PLoS Negl Trop Dis 2016;10:e0005142. doi:10.1371/journal.pntd.0005142. [8] Amanzougaghene N, Fenollar F, Sangaré AK, Sissoko MS, Doumbo OK, Raoult D, et al. Detection of bacterial pathogens including potential new species in human head lice from Mali. PLOS ONE 2017;12:e0184621. doi:10.1371/journal.pone.0184621. [9] Angelakis E, Diatta G, Abdissa A, Trape J-F, Mediannikov O, Richet H, et al. Altitude- dependent Bartonella quintana genotype C in head lice, Ethiopia. Emerg Infect Dis 2011;17:2357–9. doi:10.3201/eid1712.110453. [10] Diatta G, Mediannikov O, Sokhna C, Bassene H, Socolovschi C, Ratmanov P, et al. Prevalence of Bartonella quintana in patients with fever and head lice from rural areas of Sine-Saloum, Senegal. Am J Trop Med Hyg 2014;91:291–3. doi:10.4269/ajtmh.13-0685. [11] Murray ES, Torrey SB. Virulence of Rickettsia prowazeki for head lice. Ann N Y Acad Sci 1975;266:25–34. 75 / 285 [12] Goldberger J, Anderson JF. The Transmission of Typhus Fever, with Especial Reference to Transmission by the Head Louse (Pediculus capitis). Public Health Rep 1896-1970 1912;27:297–307. doi:10.2307/4567527. [13] Sangaré AK, Boutellis A, Drali R, Socolovschi C, Barker SC, Diatta G, et al. Detection of Bartonella quintana in African body and head lice. Am J Trop Med Hyg 2014;91:294–301. doi:10.4269/ajtmh.13-0707. [14] Kim JH, Previte DJ, Yoon KS, Murenzi E, Koehler JE, Pittendrigh BR, et al. Comparison of the proliferation and excretion of Bartonella quintana between body and head lice following oral challenge. Insect Mol Biol 2017;26:266–76. doi:10.1111/imb.12292. [15] Amanzougaghene N, Mumcuoglu KY, Fenollar F, Alfi S, Yesilyurt G, Raoult D, et al. High Ancient Genetic Diversity of Human Lice, Pediculus humanus, from Israel Reveals New Insights into the Origin of Clade B Lice. PloS One 2016;11:e0164659. doi:10.1371/journal.pone.0164659. [16] Ashfaq M, Prosser S, Nasir S, Masood M, Ratnasingham S, Hebert PDN. High diversity and rapid diversification in the head louse, Pediculus humanus (Pediculidae: Phthiraptera). Sci Rep 2015;5. doi:10.1038/srep14188. [17] Drali R, Shako J-C, Davoust B, Diatta G, Raoult D. A New Clade of African Body and Head Lice Infected by Bartonella quintana and Yersinia pestis-Democratic Republic of the Congo. Am J Trop Med Hyg 2015;93:990–3. doi:10.4269/ajtmh.14-0686. [18] Kittler R, Kayser M, Stoneking M. Molecular evolution of Pediculus humanus and the origin of clothing. Curr Biol CB 2003;13:1414–7. [19] Light JE, Allen JM, Long LM, Carter TE, Barrow L, Suren G, et al. Geographic distributions and origins of human head lice (Pediculus humanus capitis) based on mitochondrial data. J Parasitol 2008;94:1275–81. doi:10.1645/GE-1618.1. [20] Drali R, Abi-Rached L, Boutellis A, Djossou F, Barker SC, Raoult D. Host switching of human lice to new world monkeys in South America. Infect Genet Evol J Mol Epidemiol Evol Genet Infect Dis 2016;39:225–31. doi:10.1016/j.meegid.2016.02.008. [21] Raoult D, Reed DL, Dittmar K, Kirchman JJ, Rolain J-M, Guillen S, et al. Molecular identification of lice from pre-Columbian mummies. J Infect Dis 2008;197:535–43. doi:10.1086/526520. 76 / 285 [22] Boutellis A, Bitam I, Fekir K, Mana N, Raoult D. Evidence that clade A and clade B head lice live in sympatry and recombine in Algeria. Med Vet Entomol 2015;29:94–8. doi:10.1111/mve.12058. [23] Al-Shahrani SA, Alajmi RA, Ayaad TH, Al-Shahrani MA, Shaurub E-SH. Genetic diversity of the human head lice, Pediculus humanus capitis, among primary school girls in Saudi Arabia, with reference to their prevalence. Parasitol Res 2017;116:2637–43. doi:10.1007/s00436-017-5570-3. [24] Sunantaraporn S, Sanprasert V, Pengsakul T, Phumee A, Boonserm R, Tawatsin A, et al. Molecular survey of the head louse Pediculus humanus capitis in Thailand and its potential role for transmitting Acinetobacter spp. Parasit Vectors 2015;8:127. doi:10.1186/s13071-015-0742-4. [25] Candy K, Amanzougaghene N, Izri A, Brun S, Durand R, Louni M, et al. Molecular Survey of Head and Body Lice, Pediculus humanus, in France. Vector Borne Zoonotic Dis Larchmt N 2018. doi:10.1089/vbz.2017.2206. [26] Reed DL, Light JE, Allen JM, Kirchman JJ. Pair of lice lost or parasites regained: the evolutionary history of anthropoid primate lice. BMC Biol 2007;5:7. doi:10.1186/1741- 7007-5-7. [27] Tishkoff SA, Reed FA, Friedlaender FR, Ehret C, Ranciaro A, Froment A, et al. The Genetic Structure and History of Africans and African Americans. Science 2009;324:1035–44. doi:10.1126/science.1172257. [28] Ascunce MS, Fane J, Kassu G, Toloza AC, Picollo MI, González-Oliver A, et al. Mitochondrial diversity in human head louse populations across the Americas. Am J Phys Anthropol 2013;152:118–29. doi:10.1002/ajpa.22336. [29] Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: Molecular Evolutionary Genetics Analysis Version 6.0. Mol Biol Evol 2013;30:2725–9. doi:10.1093/molbev/mst197. [30] Librado P, Rozas J. DnaSP v5: a software for comprehensive analysis of DNA polymorphism data. Bioinforma Oxf Engl 2009;25:1451–2. doi:10.1093/bioinformatics/btp187. [31] Bandelt HJ, Forster P, Röhl A. Median-joining networks for inferring intraspecific phylogenies. Mol Biol Evol 1999;16:37–48. doi:10.1093/oxfordjournals.molbev.a026036. 77 / 285 [32] Boutellis A, Drali R, Rivera MA, Mumcuoglu KY, Raoult D. Evidence of sympatry of clade a and clade B head lice in a pre-Columbian Chilean mummy from Camarones. PloS One 2013;8:e76818. doi:10.1371/journal.pone.0076818. [33] Aris-Brosou S, Excoffier L. The impact of population expansion and mutation rate heterogeneity on DNA sequence polymorphism. Mol Biol Evol 1996;13:494–504. doi:10.1093/oxfordjournals.molbev.a025610. [34] Li W, Ortiz G, Fournier P-E, Gimenez G, Reed DL, Pittendrigh B, et al. Genotyping of human lice suggests multiple emergencies of body lice from local head louse populations. PLoS Negl Trop Dis 2010;4:e641. doi:10.1371/journal.pntd.0000641. 78 / 285 Figure captions Figure 1. Map showing the head lice collection site from the Amerindians of the Wayampi community living in “Trois-Sauts” Figure 2. Neighbor-joining cluster analysis of Cytb (A), Cox1 (B) and 12S (C) haplotypes of P. humanus. Bootstrap values (500 replicates) are shown above the branches. The scale bar shows K2P distances. The node for each clade with multiple haplotypes is collapsed to a vertical triangle, with the horizontal depth indicating the level of intra-clade divergence. Bracketed numbers next to each clade name indicate the number of haplotypes analyzed and the average intra-clade distance. Analyses were conducted in MEGA6. Figure 3. Cytb haplotype networks of human body and head lice. Each circle indicates a unique haplotype and variations in circle size are proportional to haplotype frequencies. Pie colors and sizes in circles represent the continents and the number of their sequence for a haplotype. Figure 4. Cox1 haplotype networks of human body and head lice. Each circle indicates a unique haplotype and variations in circle size are proportional to haplotype frequencies. Pie colors and sizes in circles represent the continents and the number of their sequence for a haplotype. Figure 5. 12S haplotype networks of human body and head lice. Each circle indicates a unique haplotype and variations in circle size are proportional to haplotype frequencies. Pie colors and sizes in circles represent the continents and the number of their sequence for a haplotype. 79 / 285 Fig. 1 80 / 285 Fig. 2 81 / 285 Fig. 3 82 / 285 Fig. 4 83 / 285 Fig. 5 84 / 285 Table 1. Primer sequences used in this study Target gene Primer Primer sequences (5′-3′) Product Tm source name size (bp) Cytochrome b cytbF GAGCGACTGTAATTACTAATC 348 56°C [34] cytbR CAACAAAATTATCCGGGTCC Cytochrome cox1F GGAGTGAGTTCGATTTTAG 828 55°C This study oxidase subunit 1 cox1R GTGCTGAGGAAAGAAAGTC 12S ribosomal 12SF CAGCACTAGCGGTCATACAT 596 56°C This study RNA 12SR AATGACGGGCGATATGTAC 85 / 285 Table 2. Analysis of genetic diversity indices and neutrality tests (Fu & Li’s D and Tajima’s D) on mitochondrial cytb, cox1 and 12S sequences n S K π h Hd Fu & Li’s D Tajima’s D Cytb all 1500 96 20.341 0.075 105 0.999 -2,878 (P <0.05) S* -0.172 (P> 0.1) NS Clade A 769 33 3.098 0.011 34 0.750 -3.751 (P <0.02) S** -2.322 (P < 0.01) S** Clade D 69 20 5.044 0.018 17 0.831 -1.425 (P> 0.1) NS -0.583 (P> 0.1) NS Clade B 200 13 3.694 0.013 9 0.789 -1.633 (P> 0.1) NS -1.357 (P> 0.1) NS Clade F 104 11 2.750 0.010 9 0.700 -1.924 (P < 0.05) S* -1.757 (P < 0.05) S* Clade C 205 16 3.744 0.014 13 0.792 -1.471 (P> 0.1) NS -1.151 (P> 0.1) NS Clade E 153 26 4.075 0.015 23 0.803 -2.747 (P < 0.05) S* -1.752 (P > 0.05) NS COI all 842 67 17.255 0.061 57 0.946 -0.065 (P > 0.1) NS 0.457 (P> 0.1) NS Clade A 443 21 3.294 0.012 17 0.769 -2.244 (P > 0.05) NS -1.866 (P < 0.05) S* Clade D 48 8 2.762 0.010 7 0.739 -0.971 (P > 0.1) NS -1.318 (P> 0.1) NS Clade B 157 12 3.697 0.013 12 0.786 -0.564 (P > 0.1) NS -0.292 (P > 0.1) NS Clade F 139 8 2.952 0.010 8 0.738 -0.473 (P > 0.1) NS -0.503 (P > 0.1) NS Clade C 25 8 3.667 0.012 6 0.782 0.457 (P > 0.1) NS 0.274 (P > 0.1) NS Clade E 30 8 3.524 0.012 7 0.781 0.321 (P > 0.1) NS 0.415 (P > 0.1) NS 12S all 442 110 26.928 0.048 49 0.964 -0.332 (P > 0.1) NS -0.130 (P > 0.1) NS Clade A 145 30 4,886 0.009 15 0.830 -2.502 (P < 0.05) S* -2.055 (P < 0.05) S* Clade D 62 23 8.821 0.016 8 0.899 0.337 (P > 0.1) NS -0.029 (P > 0.1) NS Clade B 36 3 0.271 0.001 3 0.679 -0.319 (P > 0.1) NS -1.399 (P > 0.1) NS Clade F 87 10 2.893 0.005 9 0.746 -1.612 (P > 0.1) NS -1.589 (P > 0.05) NS Clade C 69 24 7.622 0.014 10 0.885 -0.096 (P > 0.1) NS -0.483 (P > 0.1) NS Clade E 43 3 1.500 0.003 4 0.606 -0.754 (P > 0.1) NS -0.754 (P > 0.1) NS n: number of sequences; S: number of polymorphic sites; k: average number of pairwise nucleotide differences; π: nucleotide diversity; h: number of haplotypes; Hd: haplotype diversity. Tajima’s D: A negative Tajima’s D signifies an excess of low frequency polymorphisms relative to expectation. A positive Tajima’s D signifies low levels of both low and high frequency polymorphisms. Statistical significance: Not significant, P > 0.10. 86 / 285 Supporting Information Figure S1. Maximum-likelihood (ML) analysis of Cytb (A), Cox1 (B) and 12S (C) haplotypes of Pediculus humanus. Bootstrap values (500 replicates) are shown above the branches. The scale bar shows K2P distances. The node for each clade with multiple haplotypes is collapsed to a vertical triangle, with the horizontal depth indicating the level of intra-clade divergence. Bracketed numbers next to each clade name indicate the number of haplotypes analyzed and the average intra-clade distance. Analyses were conducted in MEGA6. 87 / 285 Table S1. Geographic occurrences and frequencies of cytb haplotypes of human head and body lice. Total Accessionnumber HeadBody or Africa Asia Europe Oceania America Algeria Burundi Ethiopia Kenya Madagascar Rwanda Senegal Egypt Africa South Mali DRCongo CongoBraz China Iran Japan Laos Nepal Phillippines Russia Taiwan Yemen Israel France Germany Hungary Portugal Kingdom United Australia NewZelande Papua-new-guinea Colombia Brazil Canada Ecuador Honduras Mexico Panama Peru USA Chille Argentina Niger Mongolia Pakistan Amazonia Haplotypes A1 1 1 KM579538 B A2 1 1 KM579539 B A3 1 1 KM579540 B A4 4 4 KM579541 B A5 18 21 7 17 13 91 16 24 2 1 1 45 5 27 1 9 7 69 4 10 4 38 10 17 2 1 10 1 3 1 1 9 34 1 520 KM579542 H&B A6 3 3 KM579543 B A7 1 1 KM579544 B A8 1 2 3 KM579545 B A9 1 1 KM579546 B A10 20 20 KM579547 B A11 10 10 KM579548 B A12 1 7 8 16 KM579549 H&B A13 6 5 11 KM579550 H&B A14 1 1 KM579551 H&B A16 9 9 18 KM579552 H A17 2 6 64 1 5 4 10 6 8 106 KM579553 H&B A18 2 2 KM579554 H A19 1 1 KM579555 H A45 1 1 KM579566 B A55 4 4 KX232678 H A56 1 1 KX232679 H A57 5 5 KX444540 H A58 4 4 KX444541 H A59 1 1 KX444542 H A60 5 5 MF672001 H A61 2 2 MF672002 H A62 1 1 MF672003 H A63 9 9 H A64 2 2 H A65 3 3 H A66 7 7 MH230928 H A67 2 2 MH230927 H A68 2 2 MH230926 H A69 1 1 MH230925 H B31 1 1 KX249763 H B32 1 1 KM579556 H B33 5 1 16 22 KM579557 H B34 2 2 KM579558 H B36 24 59 8 22 3 28 1 20 1 166 KM579559 H B45 1 1 KX249764 H B37 1 1 KX249776 H B38 1 1 KX249777 H B39 4 4 MF672004 H F54 89 89 H F1 5 5 H F2 1 1 H F3 2 2 H F4 1 1 H F5 1 1 H F6 1 1 H F7 3 3 H P.mjobergi 1 1 KM579537 C40 1 1 KM579561 H C41 17 79 2 98 KM579562 H C42 7 7 KM579563 H C43 1 1 KM579564 H C44 51 51 KM579565 H C51 1 1 KX249765 H C45 1 1 KX249778 H C74 32 32 KX444547 H C75 6 6 KX444548 H C76 3 3 KX444549 H C77 2 2 KX444550 H C78 1 1 KX444551 H C79 1 1 KX444552 H E39 5 14 19 KM579560 H E46 22 22 KX249780 H E47 15 15 KX249781 H E48 15 15 KY937987 H E49 2 2 KY937988 H E50 1 1 KY937989 H E51 1 1 KY937990 H E52 3 3 H E53 32 32 H E54 1 1 H E55 1 1 H E56 9 9 MF672005 H E57 3 3 MF672006 H E58 1 1 MF672007 H E59 7 7 MF672008 H E60 1 1 MF672009 H E61 1 1 MF672010 H E62 10 10 MH230921 H E63 4 4 MG759552 H E64 2 2 MG759553 H E65 1 1 MG759554 H E66 1 1 MG759555 H E67 1 1 MG759556 H D60 5 3 8 KX249766 B D61 2 2 KX249767 B D62 2 2 KX249768 B D63 3 3 KX249769 H D64 2 2 KX249770 H D65 5 11 16 KX249771 H D66 4 4 KX249772 H D67 7 7 KX249773 H D68 1 1 KX249779 H D69 3 3 KX249774 B D70 1 1 KX249775 H D71 4 4 KX444544 H D72 1 1 KX444545 H D73 2 2 KX444546 H D74 4 4 MH230924 H D75 1 1 MH230923 H D76 8 8 MH230922 H 26 18 90 7 18 13 11 91 59 70 68 177 37 6 2 10 1 6 83 15 30 1 1 89 20 144 9 10 4 32 57 16 18 120 10 7 1 12 10 1 11 84 2 1500 88 / 285 Table S2. Geographic occurrences and frequencies of cox1 haplotypes of human head and body lice. Haplotype Total Accession number or Head Body Africa Asia Europe Oceania America Algeria Burundi Ethiopia CongoDR Zimbabwe Mali Nepal Phillippines Yemen France Norway United Kingdom Australia Cook Islands Argentina Colombia Peru Ecuador Amazonia Honduras Panama USA Mexico Canada Papua-new-guinea Mongolia China A1 18 18 KM579493H A2 8412511219 195 2214 11 7 256 KM579500 H&B A3 3 2233161 1092 4 127 KM579501H A4 2 2KM579502H A5 2 2KM579503H A6 1 1KM579504H A7 22KM579505B A8 2 2KM579506H A9 3 3KM579507H A10 7 7KM579494H A11 22KM579495H A12 8 8KM579496B A13 11KM579497B A14 1 1H A15 6 6H A16 2 2B A17 33H D17 7 7KM579498B D18 2 2KM579499 H&B D19 10 10 H&B D20 4 4H D21 2 2MG759561 H&B D22 3 3MG759562B D23 20 20 B B16 16 1024 1 42 5 98 KM579508H B17 251742 KM579509H B19 33KM579511H B20 44KM579512H B21 11KM579513H B22 22KM579514H B23 11KM579515H B24 11KM579516H B25 11KM579517H B26 22KM579518H B27 11KM579519H B28 1 1KM579520H F18 10 2 12 H F29 111111 H F30 11H F31 55H F32 11H F33 55H F34 33 P.mjobergi 1 1H C31 12 12 KM579523H C32 2 2KM579524H C33 3 3MG759557H C34 1 1MG759558H C35 1 1H C36 6 6H E37 12 12 MG759559H E38 1 1MG759560H E39 1 1H E40 3 3MG759563H E41 1 1MG759564H E42 4 4MG759565H E43 8 8 MG759566 H 19 2 36 13 20 14 1 14 14 12 38 4 31 419 236 212 140 29 2 178 14 7 842 89 / 285 Table S3. Geographic occurrences and frequencies of 12S haplotypes of human head and body lice. Total number Acc. Head or Body Haplotype Africa Asia Europe Oceania America Algeria Tunisia Ethiopia Madagascar Senegal Mali CongoDR CongoBraz zimbabwe China Nepal Russia Israel France Australia USA Argentina Amazonia A1 9 6 8 11 11 2 6 1 54 H&B A2 1 6 20 3 30 H&B A3 12 6 1 19 H A4 3 3 H A5 1 3 4 B A6 1 1 H A7 2 2 H A8 2 2 H A17 6 6 H A18 2 2 H A19 11 11 H A20 4 4 B A21 3 3 B A22 1 1 H A23 3 3 H D22 13 13 H&B D23 20 20 B D24 8 8 H&B D25 3 7 10 H&B D26 3 3 H D27 2 2 B D28 1 1 B D29 5 5 H B17 12 15 6 33 H B18 1 1 H B19 2 2 H F19 70 70 H F20 7 7 H F21 2 2 H F22 1 1 H F23 1 1 H F24 1 1 H F25 1 1 H F26 1 1 H P.mjobergi 3 3 C20 24 24 H C21 5 5 H C22 3 3 H C23 6 6 H C24 2 2 H C25 7 7 H C26 6 6 H C27 1 1 H C28 11 11 H C29 4 4 H E30 16 15 31 H E31 7 7 H E32 3 3 H E33 2 2 H 24 11 41 4 22 27 17 77 20 4 20 11 11 38 12 98 2 3 442 90 / 285 Chapitre II : Epidémiologie des poux et pathogènes associés 91 / 285 Préambule L’importance des poux en santé publique humaine est surtout liée à leur capacité de transmission de maladies, plus particulièrement le pou de corps qui est le principal vecteur de R. prowazekii, B. quintana et B. recurrentis, il est aussi soupçonné dans la transmission de Y. pestis [17,18]. Le statut vectoriel du pou de tête, quant à lui, reste jusqu’à présent un sujet de controverse, bien que plusieurs études épidémiologiques et expérimentales aient montré qu’il pouvait aussi être vecteur, bien que plus faiblement comparé au pou de corps. Dans ce projet, nous avons étudié les pathogènes associés aux poux et plus particulièrement ceux associés aux poux de tête. Dans un premier temps, nous avons développé une nouvelle technique de PCR en temps réel pour l’identification moléculaire rapide des clades de poux. Par la suite, l’analyse d’une large collection de poux provenant de différents pays a mis en évidence la présence de l’ADN de plusieurs bactéries. Entre autres, l’ADN de B. quintana a été détecté chez les poux de tête du clade E du Mali, B. recurrentis chez des poux de tête du clade A des pygmées de la République du Congo. Par ailleurs, l’ADN de plusieurs autres espèces bactériennes qui n’étaient pas habituellement associées aux poux a été détecté pour la premier fois à savoir : Coxiella burnetii dans les poux de tête du clade E du Mali et des réfugiés nigériens, R. aeschlimannii et de potentiels nouvelles espèces de genre Anaplasma et Ehrlichia chez les poux de tête du clade E du Mali, ainsi que B. theileri chez les poux de tête clade A des pygmées de la République du Congo. Enfin, l’ADN de plusieurs espèces d’Acinetobacter, dont plusieurs potentielles nouvelles espèces, a été également détecté chez les poux de tête et de corps appartenant aux cinq différents clades (clades A, D, B, C et E) provenant de plusieurs pays (République du Congo, République Démocratique du Congo, France ainsi que parmi les poux des réfugiés nigériens en Algérie). 92 / 285 Article 4 : Head Lice of Pygmies Reveal the Presence of Relapsing Fever Borreliae in the Republic of Congo. Publié dans PLOS Neglected Tropical Diseases 2016; 10: e0005142 93 / 285 RESEARCH ARTICLE Head Lice of Pygmies Reveal the Presence of Relapsing Fever Borreliae in the Republic of Congo Nadia Amanzougaghene1, Jean Akiana2, Ge´or Mongo Ndombe2, Bernard Davoust1, Nardiouf Sjelin Nsana2, Henri-Joseph Parra2, Florence Fenollar1, Didier Raoult1,3*, Oleg Mediannikov1,3* 1 Unite´ de Recherche sur les Maladies Infectieuses Tropicales Emergentes (URMITE), Aix-Marseille Universite´, Marseille, France, 2 Laboratoire national de sante´ publique, Brazzaville, Re´publique du Congo, a11111 3 Campus International UCAD-IRD, Dakar, Senegal * [email protected] (OM); [email protected] (DR) Abstract OPEN ACCESS Citation: Amanzougaghene N, Akiana J, Mongo Background Ndombe G, Davoust B, Nsana NS, Parra H-J, et al. Head lice, Pediculus humanus capitis, occur in four divergent mitochondrial clades (A, B, C (2016) Head Lice of Pygmies Reveal the Presence and D), each having particular geographical distributions. Recent studies suggest that head of Relapsing Fever Borreliae in the Republic of Congo. PLoS Negl Trop Dis 10(12): e0005142. lice, as is the case of body lice, can act as a vector for louse-borne diseases. Therefore, doi:10.1371/journal.pntd.0005142 understanding the genetic diversity of lice worldwide is of critical importance to our under- Editor: Job E Lopez, Baylor College of Medicine, standing of the risk of louse-borne diseases. UNITED STATES Received: July 13, 2016 Methodology/Principal Findings Accepted: October 27, 2016 Here, we report the results of the first molecular screening of pygmies’ head lice in the Published: December 2, 2016 Republic of Congo for seven pathogens and an analysis of lice mitochondrial clades. We developed two duplex clade-specific real-time PCRs and identified three major mitochon- Copyright: © 2016 Amanzougaghene et al. This is an open access article distributed under the terms drial clades: A, C, and D indicating high diversity among the head lice studied. We identified of the Creative Commons Attribution License, the presence of a dangerous human pathogen, Borrelia recurrentis, the causative agent of which permits unrestricted use, distribution, and relapsing fever, in ten clade A head lice, which was not reported in the Republic of Congo, reproduction in any medium, provided the original author and source are credited. and B. theileri in one head louse. The results also show widespread infection among head lice with several species of Acinetobacter. A. junii was the most prevalent, followed by A. Data Availability Statement: All sequences of cytb haplotypes of Pediculus humanus, flab sequences ursingii, A. baumannii, A. johnsonii, A. schindleri, A. lwoffii, A. nosocomialis and A. towneri. of Borrelia, rpoB sequences of Acinetobacter species and Moraxellaceae are available in GenBank under accession number: KX444538- Conclusions/Significance KX444552, KX444533-KX444534, KX444507- Our study is the first to show the presence of B. recurrentis in African pygmies’ head lice in KX444532 and KX444535-KX444537, respectively. the Republic of Congo. This study is also the first to report the presence of DNAs of B. thei- Funding: The authors thank IHU Me´diterrane´e leri and several species of Acinetobacter in human head lice. Further studies are needed to Infection for financially supporting the study. The determine whether the head lice can transmit these pathogenic bacteria from person to funders had no role in study design, data collection and analysis, decision to publish, or preparation of another. the manuscript. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 1 / 18 94 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Competing Interests: The authors have declared that no competing interests exist. Author Summary Head lice, Pediculus capitis humanus, and body lice, Pediculus h. humanus, are obligatory ectoparasites that feed exclusively on human blood. Currently, the body louse is the only recognized vector of at least three deadly bacterial pathogens that have killed millions of peoples, namely: Rickettsia prowazekii, Bartonella quintana and Borrelia recurrentis, responsible for epidemic typhus, trench fever and relapsing fever, respectively. In this work, we aimed to study the genetic diversity of head lice collected from African Pygmies in the Republic of Congo and to look for louse-borne pathogens in these lice. We detected B. recurrentis in head lice belonged to clade A that is prevalent in the Republic of Congo. Our study also show, for the first time, the presence of DNAs of B. theileri and several spe- cies of Acinetobacter in human head lice. Despite several investigations into the transmis- sibility of numerous infectious agents, no conclusive evidence has demonstrated the transmission of disease by head lice. That said, we believe that pathogens detected in head lice may be an indirect tool for evaluating the risk of louse-borne diseases in humans. Introduction The head louse, Pediculus humanus capitis, and the body louse, P. h. humanus, are obligatory hematophagous parasite that thrived exclusively on human blood for thousands of years [1, 2]. The two lice are now usually considered members of a single species as opposed to separate species [3, 4], each louse lives and multiplies in a specific ecological niche: hair for head lice and clothing for body lice [5, 6]. Molecular analysis of mitochondrial genes has permitted the classification of Pediculus huma- nus into three several clades or haplogroups, referred to as A, B, and C [1, 2, 7, 8, 9]. Haplogroup A is the most common, and possesses a global distribution, including both head and body lice [1, 2, 6, 8, 9]. Clade B comprises only head lice, is confined to the New Word, Europe, Australia and was recently reported in North and South Africa [2, 6, 10, 11]. Clade C includes only head lice and is mainly found in Africa and Asia [2, 5, 9, 10]. Most recently, a novel clade D, compris- ing both head and body lice, was described in Democratic Republic of the Congo [6]. Prior research suggested that the known lice clades evolved on different lineages of Homo, similarly to those which are known to have existed 2.3 to 0.03 million years ago (MYA) [1, 11], and accordingly their geographic distribution may provide information regarding the evolu- tionary history of the lice as well as their human hosts [1, 2, 22]. Clade A lice are most likely to have emerged in Africa and to have evolved on the host linage that led to anatomically modern humans (Homo sapiens), showing the signs of a recent demographic expansion out of Africa about 100,000 years ago, first to Eurasia and subsequently to Europe, Asia, and the New World [1, 5, 12]. Haplogroup B diverged from haplogroup A between 0.7 and 1.2 MYA and may have evolved on archaic hominids, such as the Homo sapiens neanderthalensis, who spread across Europe and Asia, only becoming associated with modern humans during the period of overlap as the result of a recent host switch [1, 5, 12]. Head lice are one of the most prevalent parasitic infestations in contemporary populations, particularly in children. They often cause intense itching and, in some cases, insomnia. As a result, they represent a major economic and social concern worldwide [6, 13, 14]. Body lice, unlike head lice, are nowadays less prevalent and tend to appear mainly in indigent individuals living in poor sanitary conditions [6, 9, 13]. They do, however, present a far more serious threat to public health because they transmit at least three deadly bacterial pathogens that have killed millions of peoples, namely: Rickettsia prowazekii, Bartonella quintana, and Borrelia PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 2 / 18 95 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever recurrentis, responsible for epidemic typhus, trench fever, and relapsing fever, respectively [5, 9, 13]. Body lice are also suspected of transmitting the agent of plague, Yersinia pestis and the nosocomial pathogen, Acinetobacter baumannii [6, 15, 16]. Until recently, it was believed that head lice cannot transmit louse-borne diseases [17]. Recently, however, its status as a vector of pathogens has been brought into question, since, they have been found to carry the DNA of B. quintana, B. recurrentis, A. baumannii, and Y. pestis in natural settings [6, 18, 19, 20, 21, 22, 23]. Furthermore, experimental infections have shown that head lice may also act as a vector of louse-borne diseases [24, 25], justifying a detailed understanding of their genetic diversity and distribution worldwide. In Central Africa, studies on head lice, particularly those involving indigenous individuals, have received little prior attention. Of these indigenous populations, the African Pygmies are hunter-gatherers who live scattered in the equatorial forest. They are characterized by having a very short stature [26]. The Eastern and Western Pygmies represent the two principal groups of African Pygmies [26]. The Western group is estimated to include 55,000 individuals living in the Western Congo basin, across the countries of Cameroon, Republic of Congo, Gabon and Central African Republic, and its subgroups are identified by different names, including the Binga, Baka, Biaka and Aka or Atsua [26]. Furthermore, the detection of B. recurrentis in African lice remains limited to only a small number of countries. Currently, this bacterium is endemic in Eastern Africa (Ethiopia, Eritrea, Somalia, and Sudan) with the highest number of cases observed in Ethiopia, where it is the sev- enth most common cause of hospital admission and the fifth most common cause of death [27, 28]. Nevertheless, this borreliae has not been reported in any of the Central African coun- tries cited above. In this work, we aimed to study the genetic diversity of head lice collected from African Pygmies in the Republic of Congo and to look for louse-borne pathogens in these lice. Materials and Methods Ethics statement and louse sampling This study was approved by the Health Ministry of the Republic of Congo (000208/MSP/ CAB.15 du Ministère de la Sante´ et de la Population, 20 August 2015). All necessary permits were obtained from the individuals involved or their legal representatives in the case of chil- dren. All permissions were granted orally, because the participants are illiterate. The represen- tatives of a local Health Center and the village elders accompanied the researchers to ensure that information was correctly translated into local languages and that the villagers were will- ing to take part in the study. A total of 630 head lice samples were collected from 126 apparently healthy authochthonal individuals (pygmies) in the Republic of Congo (Congo-Brazzaville) in August 2015. The col- lections were conducted in three different villages: i) Thanry-Ipendja, where 137 lice were iso- lated from 18 people, ii) Pokola, where 163 lice were isolated from 36 people, and iii) Be´ne´- Gamboma, where 330 lice were isolated from 72 people (Fig 1). All the sampled individuals were thoroughly examined for the presence of both head and body lice. All visible head lice were removed from hair using a fine-tooth comb. Lice were then collected from the clean white tissue with forceps. No body lice were found during the examination. All the lice were preserved in 70% ethanol and transported to our laboratory in Marseille (France). DNA extraction The head lice specimens were removed from the 70% ethanol, washed three times in distilled water, and cut in half. The genomic DNA of each half louse was extracted using a DNA PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 3 / 18 96 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Fig 1. Map of head lice collection in the pygmy population from Congo-Brazzaville. doi:10.1371/journal.pntd.0005142.g001 extraction kit, QIAamp Tissue Kit (Qiagen SAS, Courtaboeuf, France) with the EZ1 apparatus following the manufacturer’s protocols. The extracted head lice DNA was assessed for quantity and quality using a Nano Drop spectrophotometer (Thermo Scientific, Wilmington, United Kingdom). The genomic DNA was stored at -20˚C under sterile conditions until the next stage of the investigation. Genotypic status of lice Determination of louse clade by real-time PCR assays. In order to identify the clades of the collected lice, we developed a real-time quantitative PCR (qPCR) method based on two duplex designed from the cytochrome b (cytb) gene. The first duplex consisted of a set of primers with FAM-and VIC- labeled probes specific to clade A and D respectively, targeting 140-bp of cytb (nucleotide position 190–329 of cytb gene). The second duplex consisted of another set of primers with FAM-and VIC- labeled probes specific to clade B and C, respectively, targeting 187-bp of the cytb gene (nucleotide position 499–685 of cytb gene). All available sequences of cytb of the four mitochondrial clades of human lice were aligned by CLUSTAL X 2.0.11 [29] and signature sites of each clade were identified. The following design was based on identified signa- ture sites and performed with Primer3 software, version 4.0 (http://frodo.wi.mit.edu/primer3/), PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 4 / 18 97 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever following the general rules described elsewhere [30]. Sequences of primers and probes are shown in Table 1. In order to confirm the specificity of the qPCRs which were developed, both duplex qPCR assays were optimized and screened for specificity and sensitivity by testing louse specimens from known clades obtained from the private frozen collection of world lice belonging to our laboratory (URMITE). All of the 630 pygmy head lice specimens were then tested in both duplex qPCR assays. The final reaction volume of 20 μl contained 5 μL of the DNA template, 10 μl of Eurogentec Probe PCR Master Mix (Eurogentec, Liège, Belgium), 0.5 mM of each primer and 0.5 mM of the FAM- and VIC labeled probes for each duplex. PCR amplification was carried out in a CFX96 Real-Time system (Bio-Rad Laboratories, Foster City, CA, USA) using the following thermal profile: one incubation step at 50˚C for two minutes and an initial denaturation step at 95˚C for three minutes, followed by 40 cycles of denaturation at 95C for 15 seconds and anneal- ing extension at 60˚C for 30 seconds. As positive controls, we used lice with known clades. Cytochrome b amplification and sequencing. For phylogenetic study, DNA samples of approximately 20% of the total number of lice collected in each village were randomly selected to ensure an equal distribution of the included lice from the three villages studied. They were subjected to standard PCR targeting a 347-bp fragment of cytb gene as previously described [31]. PCR amplification was performed in a Peltier PTC-200 model thermal cycler (MJ Research Inc., Watertown, MA, USA). PCR reactions contained 5 μl of DNA template, 2.5 μl of Tampon Buffer, 1 μl of MgCl2, 2.5 μl 2 μM of dNTP, 0.5 μl 10 μM of each primer, 0.25 μl Hotstar Taq- polymerase (Qiagen) and water to create a final reaction mixture volume of 25 μl. The thermal cycling conditions were one incubation step at 95˚C for 15 minutes, 40 cycles of one minute at 95˚C, 30 seconds at 56˚C and one minute at 72˚C followed by a final extension for five minutes at 72˚C. Negative and positive controls were included in each assay. The success of amplifica- tion was confirmed by electrophoresis on a 1.5% agarose gel. Purification of PCR products was performed using NucleoFast 96 PCR plates (Macherey-Nagel EURL, Hoerdt, France) as per the manufacturer’s instructions. The amplicons were sequenced using the Big Dye Terminator Cycle Sequencing Kit (Perkin Elmer Applied Biosystems, Foster City, CA) with an ABI auto- mated sequencer (Applied Biosystems). The electropherograms which were obtained were assembled and edited using ChromasPro software (ChromasPro 1.7, Technelysium Pty Ltd., Tewantin, Australia) and compared with those available in GenBank database by NCBI BLAST (http://blast.ncbi.nlm.nih.gov/Blast.cgi). Molecular screening for the presence of pathogen DNA The qPCR was performed to screen all lice samples using previously reported primers and probes for Borrelia spp., Bartonella spp., Acinetobacter spp., Rickettsia spp., Rickettsia prowaze- kii, Y. pestis, and Anaplasma spp. (Table 1). All qPCRs were performed using a CFX96 Real- Time system (Bio-Rad Laboratories) and the Eurogentec Master Mix Probe PCR kit (Eurogen- tec). We included the DNA of the target bacteria as positive controls and master mixtures as a negative control for each test. We considered samples to be positive when the cycle’s threshold (Ct) was lower than 35 Ct [38]. To identify the species of bacteria, all positive samples from qPCRs for Acinetobacter spp. and Borrelia spp. were further subjected to standard PCR, targeting a portion of the rpoB gene (zone1) and a portion of the flab gene, respectively, using the primers and all conditions as described previously [33, 36]. Successful amplification was confirmed via gel electrophoresis and amplicons were prepared and sequenced using similar methods as described for cytb gene for lice above. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 5 / 18 98 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Table 1. Oligonucleotide sequences of primers and probes used for real-time PCRs and conventional PCRs in this study. Target Name Primers (5’-3’) and probes Source Pediculus humanus cytochrome b Duplex A-D F_ GATGTAAATAGAGGGTGGTT This study R_ GAAATTCCTGAAAATCAAAC FAM-CATTCTTGTCTACGTTCATATTTGG-TAMRA VIC-TATTCTTGTCTACGTTCATGTTTGA-TAMRA Duplex B-C F_ TTAGAGCGMTTRTTTACCC This study R_ AYAAACACACAAAAMCTCCT FAM-GAGCTGGATAGTGATAAGGTTTAT-MGB VIC-CTTGCCGTTTATTTTGTTGGGGTTT-TAMRA Cytb F_GAGCGACTGTAATTACTAATC [31] R_CAACAAAATTATCCGGGTCC Rickettsia spp. citrate synthase (gltA) RKNDO3 F_GTGAATGAAAGATTACACTATTTAT [32] R_GTATCTTAGCAATCATTCTAATAGC FAM-CTATTATGCTTGCGGCTGTCGGTTC-TAMRA Acinetobacter spp. RNA polymerase β subunit gene rpoB F_TACTCATATACCGAAAAGAAACGG [18] R_GGYTTACCAAGRCTATACTCAAC FAM-CGCGAAGATATCGGTCTSCAAGC-TAMRA rpoB (zone1) F_TAYCGYAAAGAYTTGAAAGAAG [33] R_CMACACCYTTGTTMCCRTGA Rickettsia prowazekii rOmpB gene ompB F_AATGCTCTTGCAGCTGGTTCT [34] R_TCGAGTGCTAATATTTTTGAAGCA FAM-CGGTGGTGTTAATGCTGCGTTACAACA-TAMRA Yersinia pestis PLA F_ATG GAG CTT ATA CCG GAA AC [34] R_GCG ATA CTG GCC TGC AAG FAM-TCCCGAAAGGAGTGCGGGTAATAGG-TAMRA Borrelia spp. 16S ribosomal RNA Bor16S F_AGCCTTTAAAGCTTCGCTTGTAG [35] R_GCCTCCCGTAGGAGTCTGG FAM-CCGGCCTGAGAGGGTGAACGG-TAMRA flagellin gene flab F_GCTGAAGAGCTTGGAATGCAACC [36] R_TGATCAGTTATCATTCTAATAGCA Bartonella spp. Internal transcribed spacer 16S-23S BartoITS2 F_GATGCCGGGGAAGGTTTTC [18] R_GCCTGGGAGGACTTGAACCT FAM-GCGCGCGCTTGATAAGCGTG-TAMRA Anaplasma spp. 23S ribosomal RNA TtAna F_TGACAGCGTACCTTTTGCAT [37] R_TGGAGGACCGAACCTGTTAC FAM-GGATTAGACCCGAAACCAAG-TAMRA doi:10.1371/journal.pntd.0005142.t001 PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 6 / 18 99 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Data analysis For comparison, the head lice DNA sequences obtained in this study were combined with the 30 cytb haplotypes reported by Drali et al.[39]. We then complemented this dataset with newly available sequences in GenBank, then assigned them to haplotypes using DnaSP v5.10 [40]. Finally, we created a dataset that consisted of 51 haplotypes. These haplotypes span 41 geographic locations (countries) in five continents (S1 Table). In order to investigate the possible relationships between the haplotypes, the median-join- ing (MJ) network using the method of Bandelt was constructed with the program NET- WORK4.6 (www.fluxus-engineering.com/sharenet.htm)[41]. Phylogenetic analyses and tree reconstruction were performed using MEGA software ver- sion 6.06 [42] with 500 bootstrap replications. Results Genetic status of lice Identification of the specificity of two developed duplex qPCRs for the determination of lice clades. Two developed duplex qPCRs (A + D and B + C clades) were tested on 249 lice from the URMITE collection of those lice whose mitochondrial clades had already been identi- fied by sequencing the portion of cytb gene [31]. In total, 249/249 lice produced fluorescence curves in qPCR. The clades were correctly identified in 249/249 cases. Determination of lice clade by two duplex qPCRs. In total, 630 head lice were collected from 126 individuals living in three villages from different prefectures of Congo-Brazzaville, and all were tested by both the duplex q-PCRs to determine their clade. Our result showed that 431 (68.4%) lice belonged to clade A, 134 (21.3%) lice to clade C and only 65 (10.3%) lice to clade D. Considering the geographical regions where the lice were collected, all those collected from the villages of Pokola and Thanry-Ipendja belonged to clade A, while those collected from the village of Be´ne´-Gamboma belonged to all three clades (Clade A, C, and D). Of the 126 persons, 90 (71.41%) were mono-infested by only one clade of lice. Of these, 67 (53.17%) were only infested with lice from clade A, four (3.17%) were only infested with lice from Clade D, and 19 (15.07%) were exclusively infested with lice from Clade C. Dual infesta- tion was observed in 23 individuals (18.25%), of which eight featured both Clade A and D, seven featured both Clade A and C, and eight featured both Clade D and C. Triple infestation for all three clades was detected in 13 people (10.31%) (Table 2). Phylogenetic analysis and haplotype assignment A total of 160 head lice cytb sequences were analyzed in this work yielding 83 variable positions defining 15 different haplotypes, including 11 new ones: five from haplogroup A (35.7%), four from haplogroup D (28.5%), and six from haplogroup C (42.8%) (Table 3). These haplotypes, together with references from all the body and head lice haplogroups were used to construct a maximum-likelihood (ML) tree and a median-joining (MJ) network (Figs 2 and 3). ML and MJ analyses had similar results: all the cytb sequences were divided across the four major supported clades, represented by four connected subnetworks distinct groups as shown in the MJ network (Fig 2) corresponding to the known clades: A, D, B, and C. The 15 haplo- types in our study fell into all of the three haplogroups, A, D, and C. The haplogroup A subnetwork was star-like in structure, with the most prevalent and wide- spread haplotype being A5 (78% of locations and 45.4% of the 1,005 analyzed human lice) in the center. 24 (15%) of our cytb sequences have this A5 haplotype and are all from the village of Be´ne´-Gamboma, while a total of 64 (40%) cytb sequences (34 sequences from Thanry- PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 7 / 18 100 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Table 2. Number of pygmy individuals infested with single or multiple clades of lice from Congo- Brazzaville. Clade of lice Individual infested (n = 126) no. % Single infestation Clade A 67 53.17 Clade D 4 3.17 Clade C 19 15.07 Total 90 71.41 Multiple infestation Clade A/D 8 6.35 Clade D/C 8 6.35 Clade C/A 7 5.55 Clade A/D/C 13 10.31 Total 36 28.56 doi:10.1371/journal.pntd.0005142.t002 Ipendja and 34 sequences from Pokola villages) have the A17 haplotype, which is the second most common A-haplotype and derived from the A5-haplotype by one mutation step. The remaining five clade A sequences, four from Thanry-Ipendja and five from Pokola, defined three novel haplotypes, named here A57, A58, and A59. These three novel haplotypes derived from A17-haplotype by one mutation step. Haplogroup D, which is genetically close to A, only consists of haplotypes from Ethiopia and the Republic Democratic of Congo (RDC). The 45 (45/160) pygmy head lice sequences within clade D defined four haplotypes, of which three are novel (named here: D71, D72, D73), while the fourth haplotype possessed D65 haplotype from RDC. The clade C, representing the most divergent lineage in which two sub-clades can be defined, here referred to as sub-clade C1, which consists of head lice from Ethiopia, France and the Asian continent, and sub-clade C2, which consists of head lice from Senegal and Mali. Table 3. Haplotype frequency of pygmies’ head lice per village in Congo-Brazzaville. Haplotype Pokola Thanry-Ipendja Be´ne´-Gamboma Total Acc. no. A-5 24 24 KX444538 A-17 34 30 64 KX444539 A-57 5 5 KX444540 A-58 4 4 KX444541 A-59 1 1 KX444542 D-65 11 11 KX444543 D-71 4 4 KX444544 D-72 1 1 KX444545 D-73 1 1 KX444546 C-74 32 32 KX444547 C-75 6 6 KX444548 C-76 3 3 KX444549 C-77 2 2 KX444550 C-78 1 1 KX444551 C-79 1 1 KX444552 Total 40 34 85 160 doi:10.1371/journal.pntd.0005142.t003 PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 8 / 18 101 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Fig 2. Cytb haplotype networks of human body and head lice. Each circle indicates a unique haplotype and variations in circle size are proportional to haplotype frequencies. Pie colors and sizes in circles represent the continents and the number of their sequence for a haplotype. The length of the links between nodes is proportional to mutational differences. Haplotypes identified in the present study are in bold. doi:10.1371/journal.pntd.0005142.g002 These two subclades are separated by 12 mutations steps. Interestingly, all 45 (45/160) pygmy head lice sequences within clade C yielded six novel haplotypes, named here as C74-C79 and are parts of sub-clade C1. Molecular detection of pathogens In this study, the qPCR investigation of all 630 lice samples for Bartonella spp., Rickettsia spp., R. prowazekii, Y. pestis, and Anaplasma spp. produced no positive results. However, we obtained positive results when testing for the presence of Borrelia spp. and Acinetobacter spp. The DNA of Borrelia spp. was detected in 11/630 (1.74%) head lice collected from 7/126 (5.55%) individuals. All Borrelia-positive lice were clade A and found only in Pokola. The DNA of Acinetobacter spp. was detected in 235/630 (37.3%) head lice collected from 93/126 (73.8%) people. Of the 235 positive lice, 176 (26%) were clade A, 24 (3.8%) clade D, and 47 (7.5%) clade C. Sixty-one of these infected lice were from Pokola, forty-one from Thanry- Ipendja, and one hundred and thirty-three from Be´ne´-Gamboma. Molecular identification of Borrelia species. We succeeded in amplifying a 344-bp frag- ment of the flaB gene from all 11 lice belonging to clade A which were positive in qPCR. The comparison with the GenBank database sequences identified ten (10/11) of the obtained sequences as B. recurrentis with 100% similarity, and the one remaining sequence was identi- fied as B. theileri with 99% identity. The phylogenetic position of these Borrelia is shown in Fig 4. The sequences of these two Borrelia were deposited in the GenBank under the accession number: KX444533- KX444534. Molecular identification of Acinetobacter species. We succeeded in amplifying a frag- ment of the rpoB gene in 202 of the 235 samples that were positive in qPCR for Acinetobacter spp. The comparison of the nucleotide sequences with the GenBank database sequences revealed that only 144/202 (71.3%) sequences match eight species of Acinetobacter sharing 98–100% simi- larity, which are, in order of decreasing frequency: Acinetobacter junii (37/202; 18.31%), PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 9 / 18 102 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Fig 3. Maximum-likelihood phylogram of Pediculus humanus haplotypes based on partial 272-bp cytb gene with Pediculus schaeffi and Pthirus pubis as outgroups. doi:10.1371/journal.pntd.0005142.g003 Acinetobacter ursingii (29/202; 14.35%), Acinetobacter baumannii (22/202; 10.89%), Acinetobac- ter johnsonii (19/202; 9.40%), Acinetobacter schindleri (17/202; 8.41%), Acinetobacter lwoffii (9/ 202; 4.45%), Acinetobacter nosocomialis (7/202; 3.18%), and Acinetobacter towneri (4/202; 1.98%). The distribution of species according clade of lice and collection site are presented in PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 10 / 18 103 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Fig 4. Maximum-likelihood phylogenetic tree based on 340-bp fragment flaB gene of the Borrelia species. doi:10.1371/journal.pntd.0005142.g004 Table 4. The other 52/202 (25.74%) sequences also rated resembled Acinetobacter but were of poor quality, which is assumed to be due to co-infection with several Acinetobacter species. Six of the 202 (2.97%) remaining sequences revealed 76% identity with the sequence of Moraxella osloensis (accession no. AP017381). It may represent the DNA of an as yet uniso- lated and undescribed bacterial species of Pseudomonadales. The phylogenetic tree demon- strated that all Acinetobacter species were classified in the same group as the reference sequence strain and showed that all the Moraxellaceae bacterium were classified in the same group as the Moraxella species but formed a separate branch on the phylogenetic tree (Fig 5). The partial rpoB sequences of the Acinetobacter species and the Moraxellaceae species obtained in this study were deposited in the GenBank under the accession number KX444507-KX444532 and KX444535-KX444537, respectively. Discussion Here, we report the first molecular data on human head lice, P. h. capitis, infesting the pygmy population in the Republic of Congo in Western Africa. In this study, we established and eval- uated for the first time, qPCR assay based on two duplex designed from the cytb gene, which is very well established in the study of lice, in order to identify all known clades of P. humanus. The assay adopted herein proved itself to be fast, specific, sensitive and fully compatible when routinely analyzing large collections of lice specimens. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 11 / 18 104 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Table 4. Detection of head lice clades and pathogens in the pygmy population in Congo-Brazzaville. Villages Sample no. Acinetobacter species Borrelia species Moraxellaceae bacterium Infection (%) Infection rate no. Species identification Infection rate no. Species rate no. (%) (%) (%) identification Be´ne´-Gamboma Person 72 (57.1%) 53 1 Head 330 (52.3%) 133 2 lice Clade A 131 62 AJ, AU, AJn, AB, AS, 2 AL Clade D 65 24 AJ, AU, AJn, AB, AN, - AS, AL Clade C 134 47 AJ, AU, AJn, AB, AN, - AS, AT Thanry-Ipendja Person 18 (14.3%) 12 1 Head 137 (21.7%) 41 1 lice Clade A 137 41 AJ, AU, AJn, AB, AN 1 Pokola Person 36 (28.6%) 28 7 2 Head 163 (25.9%) 61 11 3 lice Clade A 163 61 AJ, AU, AJn, AB, AS, 11 BR (n = 10), BT 3 AL (n = 1) Total Person 126 93 (73.8%) 7 4 Head 630 235 (37.3%) 11 6 lice Clade A 431 (68.4%) 164 (26%) AJ, AU, AJn, AB, AN, 11 BR, BT 6 AS, AL Clade D 65 (10.3%) 24 (3.8%) AJ, AU, AJn, AB, AN, --- AS, AL Clade C 134 (21.3%) 47 (7.5%) AJ, AU, AJn, AB, AN, --- AS, AT AJ: Acinetobacter junii; AU: A. ursingii; AJn: A.johnsonii; AB: A.baumannii; AN: A. nosocomialis; AS: A. schandleri; AL: A. lwoffii; AT: A. towneri. BR: Borrelia recurrentis; BT: B. theileri. doi:10.1371/journal.pntd.0005142.t004 The mtDNA analysis of 630 head lice, collected from 126 pygmies, showed the presence of three major mitochondrial haplogroups: A, C and D, indicating high mtDNA diversity among the head lice studied. Haplogroup A was the most prevalent (56%) followed by haplogroup C (5%). The data confirm that clade A has worldwide distribution, as reported by others [6, 8, 9, 10]. Previous studies reported that clade C is limited to Nepal and Thailand [1, 5, 23], Ethiopia, Senegal and Mali [5, 9, 18, 22]; this is the first report of clade C which has been found in the Republic of Congo. The remaining samples (10.3%) were from new haplogroup D, which is known only to exist in Democratic Republic of the Congo and Ethiopia [2, 6]. In addition to inter-haplogroup diversity, P. humanus also presents intra-haplogroup diversity, illustrated by many distinct A, B and C haplotypes [2, 12, 39]. These results are supported by our finding, that, of the 160 head lice cytb sequences analysed, 15 different haplotypes were identified, of which 11 were novel. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 12 / 18 105 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Fig 5. Maximum-likelihood phylogenetic tree based on 440-bp fragment rpoB gene of the Acinetobacter species and Moraxellaceae species, while Pseudomonas was used as an out group. doi:10.1371/journal.pntd.0005142.g005 B. recurrentis is the known causative agent of relapsing fever which, if untreated, can be fatal in up to 40% of patients [13, 43, 44]. It has long been established that body lice are the main vector for this bacterial pathogen [13, 27]. In the present study, the DNA of B. recurrentis was detected in 10/630 (1.58%) head lice belonging to clade A collected from 6/126 (4.76%) individuals. Specifically, all positives cases were only found in Pokola, suggesting that a small, PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 13 / 18 106 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever unnoticed outbreak may have occurred in the population in this area. This is the second report of the presence of B. recurrentis DNA in human head lice. Recently, this bacterium was also detected in 23% of head lice clade C from patients with louse-borne relapsing fever in Ethiopia and, because these patients were also infested with body lice, the authors hypothesize that head lice might be contaminated by blood that is infected with B. recurrentis [21]. In this study, the discovery of B. recurrentis in the clade A head lice, the same clade that includes body lice, and the absence of body lice may support the hypothesis that B. recurrentis may be transmitted by clade A head lice. Nevertheless, evidence for the presence of the DNA of this bacterium in head lice by PCR cannot distinguish between transient infections, accidentally acquire the pathogen from the blood of infected individuals, and those established in a competent vector, maintain and trans- mit the pathogen. Further studies are needed to determine whether the head louse can act as a vector of B. recurrentis. Interestingly, one of the Borrelia-positive lice was identified as B. theileri. This is the first report of the presence of the DNA of this species in human head lice. B. theileri is a spirochete that causes borreliosis in cattle, a relapsing fever-like illness, transmitted by hard ticks, such as Rhipicephalus (Boophilus)[45]. This infection can be considered as being rediscovered, appears to exist in regions where diagnostic ability is limited and its impact on livestock is largely unex- plored [45]. In this study, two hypotheses can arise from the detection of B. theileri in human head lice. The first one is that the presence of this bacterium results from environmental and/or labora- tory contaminations. This hypothesis is hardly possible, because, our work was carried out in a laboratory where B. theileri had never been worked on, nor had B. theileri DNA been extracted. Indeed, each PCR assay was systematically validated by the presence of positive and negative controls. Moreover, our collection contains lice only and didn’t contain another specimens like ticks that could be an important source of environmental contamination. The second hypothesis is that, as head lice feed only on human blood [5], the acquired infection would be from the blood of patients with ongoing bacteremia. Although, humans infected with this spirochete have not been described in the literature, the transmission of this patho- gen to humans may not be ruled out. Moreover, the sequence generated in this study was more similar by flaB sequence comparison to those reported from Ornithodoros sp. soft tick (GenBank KP191621) collected from cave in Israel, than, those reported from Rhipicephalus hard tick (GenBank KF569936) from Mali, as shown in the phylogenetic tree (Fig 4). Ornitho- doros ticks can feed from multiple warm-blooded vertebrates, including humans, and are known to transmit several species of Borrelia to humans [27, 43], thus taking in consideration that the epidemiology of B. theileri is not yet completely discovered, hypothetically it may be transmitted to humans. Finally, if our hypothesis of B. theileri bacteremia in persons harboring head lice is true, this may merely reflect ‘accidental spill-over’ from animal hosts infection, such phenomena has already been described in the literature, with the finding of the DNA of B. duttonii, the species that is only know to infect ticks and humans, in chickens and swine living close to their human owners [43]. Findings from this study also show widespread infection of head lice with several species of Acinetobacter. In total, eight Acinetobacter species were detected in 144 samples; A. junii was the most prevalent, followed by A. ursingii, A. baumannii, A. johnsonii, A. schindleri, A. lwoffii, A. nosocomialis and A. towneri. The DNA of A. towneri was only found in clade C head lice, the DNA of A. lwoffii was only found in clades A and D, while the DNA of the remaining spe- cies was found in all three clades A, D and C. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 14 / 18 107 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Previous studies demonstrated that A. baumannii is the most commonly found species in body and head lice [23], as shown by its detection in 21% of body lice collected worldwide [15], in 33% of head lice collected from Parisian elementary school children, belonging to the clade A [19] and in 71% body and 47% head lice collected from healthy individuals from Ethi- opia [20]. Another study, performed in head lice samples collected from elementary school children in Thailand, showed the presence of the DNA of three Acinetobacter species in 3.62% head lice belonging to both clade A and C. The Acinetobacter species identified were A. bau- mannii, A. schindleri and A. radioresistens [23]. When comparing the panel of Acinetobacter species found in all these studies with our findings, A. radioresistens was the only species that we did not identify in our head lice specimens. Conversely, our sampling showed, for the first time, the presence of the DNA of A. junii, A. ursingii, A. johnsonii, A. lwoffii, A. nosocomialis and A. towneri in human head lice, but further study is needed to determine the significance of this finding. Furthermore, it is still unknown how these lice acquire their Acinetobacter infections. Some authors have argued that the infection could occur after the ingestion of infected blood meal from individuals with ongoing bacteremia, or may possibly be derived from superficial con- tamination through human skin while feeding [15]. An experimental study showed that the human body louse, feeding on bacteremic rabbits, is able to acquire and maintain a persistent life-long infection with A. baumannii and A. lwoffii [46]. Furthermore, another study per- formed a comparison between two sequenced genomes of A. baumannii and showed that the A. baumannii SDF strain, isolated from a human body louse, had several hundred insertion sequence elements which have played a crucial role in its genome reduction (gene disruptions and simple DNA loss) compared to the human multidrug-resistant A. baumannii AYE strain, and also been shown to have low catabolic capacities, suggesting the specific adaptation of this strain to the louse environment [47]. However, Acinetobacter species are widespread in nature (water, soil, living organisms, and the skin of patients and healthy subjects) [47], and because the frequency of with which these species associate with the skin of pygmy population is unknown, it is not possible to rule out the infection of lice by external contamination. Clinically, A. baumannii is known to be a major cause of nosocomial infections in humans and it is an increasing public health concern due to the increasing resistance to antibiotic treatment which has been identified worldwide [47]. Other Acinetobacter species include A. lwoffii and A. junii are also often identified as the cause of infection in humans [48]. However, it still not clear whether these Acinetobacter strains present in lice are the same as those that are responsible for human infections [20]. Conclusions In conclusion, the qPCR adopted in this study proved to be a fast, sensitive and specific tool that is fully compatible when routinely analyzing a large collections of lice specimens. Our results showed the presence of three major mitochondrial haplogroups: A, C and D, indicating high mtDNA diversity among the pygmy head lice studied. We identified the presence of a dangerous human pathogen, B. recurrentis, the causative agent of relapsing fever, in ten clade A head lice, which had not previously been reported in the Republic of Congo. Findings from this study also show the widespread infection of head lice with several species of Acinetobacter. Despite several investigations into the transmissibility of numerous infectious agents, no conclusive evidence has demonstrated the transmission of disease by head lice. That said, we believe that pathogens detected in head lice may be an indirect tool for evaluating the risk of louse-borne diseases in humans. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 15 / 18 108 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever Supporting Information S1 Checklist. STROBE Checklist. (DOC) S1 Table. Geographic occurrences and frequencies of cytb haplotypes of human head and body lice. Haplotypes highlighted in blue are the newly identified haplotypes from sequences available in GenBank. (XLSX) Acknowledgments The authors thank all the villagers who participated in the study and the staff of the local Health Centers for their good will. Author Contributions Conceived and designed the experiments: OM FF DR JA. Performed the experiments: NA FF DR OM. Analyzed the data: NA BD FF DR OM. Contributed reagents/materials/analysis tools: JA GMN NSN BD OM FF DR. Wrote the paper: NA OM FF DR JA BD HJP GMN NSN. References 1. Reed DL, Smith VS, Hammond SL, Rogers AR, Clayton DH. Genetic analysis of lice supports direct contact between modern and archaic humans. PLoS Biol. 2004; 2:e340. doi: 10.1371/journal.pbio. 0020340 PMID: 15502871 2. Light JE, Allen JM, Long LM, Carter TE, Barrow L, Suren G, et al. Geographic distributions and origins of human head lice (Pediculus humanus capitis) based on mitochondrial data. J Parasitol. 2008; 94:1275–1281. doi: 10.1645/GE-1618.1 PMID: 18576877 3. Xiong H, Campelo D, Boutellis A et al. SNPs in Entire Mitochondrial Genome Sequences (approxi- mately 15.4 kb) and cox1 Sequences (approximately 486bp) Resolve Body and Head Lice From Doubly Infected People From Ethiopia, China, Nepal, and Iran But Not France. J Med Entomol. 2014a; 51:1199–1207. 4. Tovar-Corona JM, Castillo-Morales A, Chen L et al. Alternative Splice in Alternative Lice. Molecular Biol- ogy and Evolution. 2015; 32 (10):2749–2759. doi: 10.1093/molbev/msv151 PMID: 26169943 5. Boutellis A, Abi-Rached L, Raoult D. The origin and distribution of human lice in the world. Infect Genet Evol. 2014; 23: 209–217. doi: 10.1016/j.meegid.2014.01.017 PMID: 24524985 6. Drali R, Shako JC, Davoust B, Diatta G, Raoult D. A new clade of African body and head lice infected by Bartonella quintana and Yersinia pestis—Democratic Republic of Congo. Am. J. Trop. Med. Hyg. 2015; 93: 990–993. doi: 10.4269/ajtmh.14-0686 PMID: 26392158 7. Kittler R, Kayser M, Stoneking M. Molecular evolution of Pediculus humanus and the origin of clothing. Curr Biol. 2003; 13: 1414–1417. PMID: 12932325 8. Raoult D, Reed DL, Dittmar K, Kirchman JJ, Rolain JM, et al. Molecular identification of lice from pre- Columbian mummies. J Infect Dis. 2008; 197: 535–543. doi: 10.1086/526520 PMID: 18254682 9. Xiong H, Campelo D, Pollack RJ et al. Second-generation sequencing of entire mitochondrial coding- regions (approximately 15.4 kb) holds promise for study of the phylogeny and taxonomy of human body lice and head lice. Med Vet Entomol. 2014b; 28 Suppl 1:40–50. 10. Boutellis A, Bitam I, Fekir K, Mana N, Raoult D. Evidence that clade A and clade B head lice live in sym- patry and recombine in Algeria. Med Vet Entomol. 2015; 29: 94–98. doi: 10.1111/mve.12058 PMID: 25346378 PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 16 / 18 109 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever 11. Ashfaq M, Prosser S, Nasir S, Masood M, Ratnasingham S, Hebert PDN. High diversity and rapid diver- sification in the head louse, Pediculus humanus (Pediculidae: Phthiraptera). Scientific Reports. 2015; 5:14188. doi: 10.1038/srep14188 PMID: 26373806 12. Ascunce MS, Fane J, Kassu G, Toloza AC, Picollo MI, Gonzalez-Oliver A, et al. Mitochondrial diversity in human head louse populations across the Americas. Am J Physiol Anthropol. 2013; 152 (1):118– 129. 13. Raoult D, Roux V. The body louse as a vector of reemerging human diseases. Clin Infect Dis. 1999; 29: 888–911. doi: 10.1086/520454 PMID: 10589908 14. Veracx A, Raoult D. Biology and genetics of human head and body lice. Trends Parasitol. 2012; 28: 563–571. doi: 10.1016/j.pt.2012.09.003 PMID: 23069652 15. La Scola B, Raoult D. Acinetobacter baumannii in human body louse. Emerg Infect Dis. 2004; 10:1671–3. doi: 10.3201/eid1009.040242 PMID: 15498175 16. Houhamdi L, Lepidi H, Drancourt M, Raoult D. Experimental model to evaluate the human body louse as a vector of plague. J Infect Dis. 2006; 194: 1589–1596. doi: 10.1086/508995 PMID: 17083045 17. Robinson D, Leo N, Prociv P, Barker SC. Potential role of head lice, Pediculus humanus capitis, as vec- tors of Rickettsia prowazekii. Parasitology Research. 2003; 90:209–211. doi: 10.1007/s00436-003- 0842-5 PMID: 12783309 18. Angelakis E, Diatta G, Abdissa A, Trape JF, Mediannikov O, Richet H, et al. Altitude-dependent Barto- nella quintana genotype C in head lice, Ethiopia. Emerg Infect Dis. 2011; 17: 2357–2359. doi: 10.3201/ eid1712.110453 PMID: 22172306 19. Bouvresse S, Socolovschi C, Berdjane Z, Durand R, Izri A, Raoult D, et al. No evidence of Bartonella quintana but detection of Acinetobacter baumannii in head lice from elementary school children in Paris. Comp Immunol Microbiol Infect Dis. 2011; 34: 475–477. doi: 10.1016/j.cimid.2011.08.007 PMID: 21974965 20. Kempf M, Abdissa A, Diatta G, Trape JF, Angelakis E, Mediannikov O, et al. Detection of Acinetobacter baumannii in human head and body lice from Ethiopia and identification of new genotypes. Int J Infect Dis. 2012; 16: e680–e683. doi: 10.1016/j.ijid.2012.05.1024 PMID: 22771379 21. Boutellis A, Mediannikov O, Bilcha KD, Ali J, Campelo D, Barker SC, et al. Borrelia recurrentis in head lice, Ethiopia. Emerg Infect Dis. 2013; 19: 796–798. doi: 10.3201/eid1905.121480 PMID: 23648147 22. Sangare AK, Boutellis A, Drali R, Socolovschi C, Barker SC, Diatta G, et al. Detection of Bartonella quintana in African body and head lice. Am J Trop Med Hyg. 2014; 91: 294–301. doi: 10.4269/ajtmh. 13-0707 PMID: 24935950 23. Sunantaraporn S, Sanprasert V, Pengsakul Th, Phumee A, Boonserm, Tawatsin A, et al. Molecular sur- vey of the head louse Pediculus humanus capitis in Thailand and its potential role for transmitting Acine- tobacter spp. Parasit Vectors. 2015; 8: 127. doi: 10.1186/s13071-015-0742-4 PMID: 25889008 24. Goldberger J, Anderson JF. The transmission of Typhus fever, with especial reference to transmission by the head louse (Pediculus capitis). Public Health Reports (1896–1970). 1912; 27: 297–307. 25. Murray ES, Torrey SB. Virulence of Rickettsia prowazekii for head lice. Ann N Y Acad Sci. 1975; 266: 25–34. PMID: 829471 26. Batini C, Lopes J, Behar DM, Calafell F, Jorde LB, van der Veen L, et al. Insights into the demographic history of African Pygmies from complete mitochondrial genomes. Mol Biol Evol. 2011; 28: 1099–1110. doi: 10.1093/molbev/msq294 PMID: 21041797 27. Cutler SJ. Relapsing fever—a forgotten disease revealed. J Appl Microbiol. 2009; 108:1115–1122. doi: 10.1111/j.1365-2672.2009.04598.x PMID: 19886891 28. Antinori S, Mediannikov O, Corbellino M, Grande R, Parravicini C, Bestetti G, et al. Louse-Borne Relapsing Fever (Borrelia recurrentis) in a Somali Refugee Arriving in Italy: A Re-emerging Infection in Europe?. PLoS Neglected Tropical Diseases. 2016; 10(5):e0004522. http://doi.org/10.1371/journal. pntd.0004522. doi: 10.1371/journal.pntd.0004522 PMID: 27149282 29. Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, McWilliam H, et al. Clustal W and Clustal X version 2.0. Bioinformatics. 2007; 23: 2947–2948. doi: 10.1093/bioinformatics/btm404 PMID: 17846036 30. Thornton B, Basu C. Real-time PCR (qPCR) primer design using free online software. Biochem Mol Biol Educ. 2011; 39: 145–154. doi: 10.1002/bmb.20461 PMID: 21445907 31. Li W, Ortiz G, Fournier PE, Gimenez G, Reed DL, Pittendrigh B. et al. Genotyping of human lice sug- gests multiple emergencies of body lice from local head louse populations. PLoS Negl Trop Dis. 2010; 4:e641. doi: 10.1371/journal.pntd.0000641 PMID: 20351779 32. Rolain JM, Sthul L, Maurin M, Raoult D. Evaluation of antibiotic susceptibilities of three Rickettsial spe- cies including Rickettsia felis by a quantitative PCR DNA Assay. Antimicrob Agents Chemother. 2002; 46: 2747–51. doi: 10.1128/AAC.46.9.2747-2751.2002 PMID: 12183224 PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 17 / 18 110 / 285 Head Lice of Pygmies Reveal the Presence of Relapsing Fever 33. La Scola B, Gundi VA, Khamis A, Raoult D. Sequencing of the rpoB gene and flanking spacers for molecular identification of Acinetobacter species. JCl in Microbio. 2006; l44: 827–832. 34. Nguyen-Hieu T, Aboudharam G, Signoli M, Rigeade C, Drancourt M, Raoult D Evidence of a louse- borne outbreak involving typhus in Douai, 1710–1712 during the war of Spanish succession. PLoS One. 2010; 5: e15405. doi: 10.1371/journal.pone.0015405 PMID: 21060879 35. Parola P, Diatta G, Socolovschi C, Mediannikov O, Tall A, Bassene H, et al. Tick-borne relapsing fever borreliosis, rural Senegal. Emerg Infect Dis. 2011; 17:883–5 doi: 10.3201/eid1705.100573 PMID: 21529402 36. Vial L, Diatta G, Tall A, Ba el H, Bouganali H, Durand P, et al. Incidence of tick-borne relapsing fever in West Africa: longitudinal study. Lancet. 2006; 368(9529): 37–43. doi: 10.1016/S0140-6736(06)68968- X PMID: 16815378 37. Dahmani M, Bernard D, Mohamed SB, Fenollar F, Raoult D, Mediannikov O. Development of a new PCR-based assay to detect Anaplasmataceae and the first report of Anaplasma phagocytophilum and Anaplasma platys in cattle from Algeria. Comp Immunol Microbiol Infect Dis. 2015; 39: 39–45. doi: 10. 1016/j.cimid.2015.02.002 PMID: 25748051 38. Sokhna C, Mediannikov O, Fenollar F, Bassene H, Diatta G, Tall A, et al. Point-of-care laboratory of pathogen diagnosis in rural Senegal. PLoS Negl Trop Dis. 2013; 7 (1): e1999. doi: 10.1371/journal. pntd.0001999 PMID: 23350001 39. Drali R, Abi-Rached L, Boutellis A, Djossou F, Barker SC, Raoult D. Host switching of human lice to new world monkeys in South America. Infect Genet Evol. 2016; 39: 225–231. doi: 10.1016/j.meegid. 2016.02.008 PMID: 26867815 40. Librado P, Rozas J. DnaSP v5: a software for comprehensive analysis of DNA polymorphism data. Bio- informatics. 2009; 25: 1451–1452. doi: 10.1093/bioinformatics/btp187 PMID: 19346325 41. Bandelt HJ, Forster P, Rohl A. Median-joining networks for inferring intraspecific phylogenies. Mol Biol Evol. 1999; 16: 37–48. PMID: 10331250 42. Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: molecular evolutionary genetics analy- sis version 6.0. Mol Biol Evol. 2013; 30: 2725–2729. doi: 10.1093/molbev/mst197 PMID: 24132122 43. Cutler SJ, Abdissa A, Trape JF. New concepts for the old challenge of African relapsing fever borrelio- sis. Clin Microbiol Infect. 2009; 15:400–406. doi: 10.1111/j.1469-0691.2009.02819.x PMID: 19489922 44. Grosskinsky S, Schott M, Brenner C et al. Borrelia recurrentis employs a novel multifunctional surface protein with anti-complement, anti-opsonic and invasive potential to escape innate immunity. PLoS One. 2009; 4:e4858. doi: 10.1371/journal.pone.0004858 PMID: 19308255 45. Cutler SJ, Ruzic-Sabljic E, Potkonjak A. Emerging borreliae—Expanding beyond Lyme borreliosis. Molecular and Cellular Probes. 2016. 46. Houhamdi L and Raoult D. Experimental infection of human body lice with Acinetobacter baumannii. Am.J. Trop. Med. Hyg. 2006; 74: 526–531. PMID: 16606978 47. Vallenet D, Nordmann P, Barbe V, Poirel L, Mangenot S, et al. Comparative Analysis of Acinetobacters: Three Genomes for Three Life styles. PLoS ONE. 2008; 3(3): e1805. doi: 10.1371/journal.pone. 0001805 PMID: 18350144 48. Tayabali AF, Nguyen KC, Shwed PS, Crosthwait J, Coleman G, Seligy VL. Comparison of the Virulence Potential of Acinetobacter Strains from Clinical and Environmental Sources. PLoS ONE. 2012; 7(5): e37024. doi: 10.1371/journal.pone.0037024 PMID: 22655033 PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0005142 December 2, 2016 18 / 18 111 / 285 S1 Table. Geographic occurrences and frequencies of cytb haplotypes of human head and body lice. Haplotypes highlighted in blue are the newly identified haplotypes from sequences available in GenBank 112 / 285 Article 5 : Detection of several emerging bacterial pathogens in human head lice from Mali Publié dans PloS One. 2017; 12:e0184621 113 / 285 RESEARCH ARTICLE Detection of bacterial pathogens including potential new species in human head lice from Mali Nadia Amanzougaghene1, Florence Fenollar1, Abdoul Karim Sangare´2, Mahamadou S. Sissoko2, Ogobara K. Doumbo2, Didier Raoult1,3*, Oleg Mediannikov1,3* 1 Aix Marseille Univ, CNRS, IRD, INSERM, AP-HM, URMITE, IHU - Me´diterrane´e Infection, Marseille, France, 2 University of Bamako, Epidemiology Department of Parasitic Diseases, Faculty of Medicine and Odonto-Stomatology, Faculty of Pharmacy (MRTC/DEAP/FMOS-FAPH), Bamako, Mali, 3 Campus a1111111111 International UCAD-IRD, Dakar, Senegal a1111111111 a1111111111 * [email protected] (OM); [email protected] (DR) a1111111111 a1111111111 Abstract In poor African countries, where no medical and biological facilities are available, the identifi- cation of potential emerging pathogens of concern at an early stage is challenging. Head OPEN ACCESS lice, Pediculus humanus capitis, have a short life, feed only on human blood and do not Citation: Amanzougaghene N, Fenollar F, Sangare´ AK, Sissoko MS, Doumbo OK, Raoult D, et al. transmit pathogens to their progeny. They are, therefore, a perfect tool for the xenodiagno- (2017) Detection of bacterial pathogens including sis of current or recent human infection. This study assessed the occurrence of bacterial potential new species in human head lice from pathogens from head lice collected in two rural villages from Mali, where a high frequency of Mali. PLoS ONE 12(9): e0184621. https://doi.org/ head lice infestation had previously been reported, using molecular methods. Results show 10.1371/journal.pone.0184621 that all 600 head lice, collected from 117 individuals, belonged to clade E, specific to West Editor: Feng Gao, Tianjin University, CHINA Africa. Bartonella quintana, the causative agent of trench fever, was identified in three of the Received: May 24, 2017 600 (0.5%) head lice studied. Our study also shows, for the first time, the presence of the Accepted: August 28, 2017 DNA of two pathogenic bacteria, namely Coxiella burnetii (5.1%) and Rickettsia aeschliman- Published: September 20, 2017 nii (0.6%), detected in human head lice, as well as the DNA of potential new species from the Anaplasma and Ehrlichia genera of unknown pathogenicity. The finding of several Copyright: © 2017 Amanzougaghene et al. This is an open access article distributed under the terms Malian head lice infected with B. quintana, C. burnetii, R. aeschlimannii, Anaplasma and of the Creative Commons Attribution License, Ehrlichia is alarming and highlights the need for active survey programs to define the public which permits unrestricted use, distribution, and health consequences of the detection of these emerging bacterial pathogens in human reproduction in any medium, provided the original head lice. author and source are credited. Data Availability Statement: All sequences of cytb haplotypes of Pediculus humanus, gltA sequences of R. aeschlimannii, groEl sequences of Ehrlichia and rpoB sequences of Anaplasma were deposited in the GenBank under accession number: Introduction KY937987-KY937990, KY937991- KY937992, KY937978- KY937986, respectively. Humans are parasitized by three different types of sucking lice (Anoplura): the head louse, the body louse and the crab (pubic) louse, each of them colonizing a specific region of the body Funding: This study was funded by the IHU (head, body and pubic area, respectively) [1,2]. Two of these types are of great concern to pub- Me´diterrane´e Infection (http://en.mediterranee- infection.com/). The funders had no role in study lic health and are now believed to be members of a single species, Pediculus humanus, which design, data collection and analysis, decision to appears in two ecotypes P. h. capitis (known as the head louse) and P. h. humanus (also known publish, or preparation of the manuscript. as the body or clothing louse) [3,4]. Both ecotypes have the same life cycle and feed exclusively PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 1 / 18 114 / 285 Detection of bacterial pathogens in head lice from Mali Competing interests: The authors have declared on human blood. They nevertheless occupy distinct ecological niches and have distinctly dif- that no competing interests exist. ferent feeding patterns [1,3,4]. Head lice live exclusively in the scalp region of humans, where the females lay eggs (nits) at the base of hair shafts [1,3]. They are prevalent worldwide, partic- ularly in school-aged children, regardless of hygiene conditions and can cause very intense pruritus that may lead to high irritation and even wound infection [4–6]. In contrast, body lice feed on the body regions of humans and the females secure their eggs to clothing [1,3]. They were also very common in the past, but are more rarely encountered in modern times and tend to be restricted to precarious populations living in poor sanitary conditions, such as the homeless, war refugees and the prison population [4,5]. Head lice have been considered to be the ancestral lineage from which body lice have rela- tively recently emerged and probably on multiple occasions [3,5,7,8]. However, this claim is not supported by genomic analysis, as only one nuclear genetic marker has been identified based on variations in the PHUM540560 gene, which encodes a hypothetical 69-amino acids protein of unknown function that can unambiguously distinguish head from body lice once they are removed from their habitat. In body lice, this gene is present and expressed, whereas it is present but not expressed in head lice (deleted) [9]. In contrast, genetic studies based on mitochondrial DNA (mtDNA) appear to separate head lice into five divergent mitochondrial clades (A, B, C, D and E) and place body lice only in two clades, A and D [3,10,11], together with head lice. Clade A is the most common and spread worldwide, while clade D has to this point only been found in Africa [12]. Clade B is found in America, Europe, Australia, North and South Africa, and was most recently reported in Israel on head lice remains dating from approximately 2,000 years ago [11,13]. Clade C is found in Ethiopia, the Republic of Congo, the Asian continent and, recently, in France [3,12]. Lastly, the fifth clade, which has previously been described as sub-clade within clade C [12,14], and which has been only recently classified as a separate new clade is referred here as clade E [11]. This clade consists of head lice from West Africa (Senegal and Mali) [11,13]. All these data support the hypothesis that all current human lice travelled with archaic hominids (and slaves) from Africa. Human body lice are the main vectors of three serious human pathogens: Rickettsia prowa- zekii (the causative agent of epidemic typhus), Bartonella quintana (trench fever) and Borrelia recurrentis (relapsing fever) [1,5]. There are natural and experimental observations that body lice can also transmit Yersinia pestis, the causative agent of plague, and that they may be the pandemic vectors of this agent [15–17]. Some other widespread pathogenic bacteria, such as Serratia marcescens, Acinetobacter baumannii and A. lwoffii, have been detected in human body lice with the assumption that lice can probably also transmit these agents to humans [4,18,19]. Early field observation in East Africa showed that human lice collected from a place where an epidemic of Q fever occurred three months previously, contained its agent, C. burne- tii. Bacterial strains were recovered from these lice using guinea pigs [20]. Experimentally infected body lice are also capable of transmitting R. typhi (the causative agent of endemic or murine typhus), R. rickettsii (Rocky Mountain spotted fever) and R. conorii (Mediterranean spotted fever, Indian tick typhus) to rabbits [21,22]. Although body lice are much more potent vectors of pathogens than head lice, perhaps due to the size of the blood meal ingested (body lice ingest a larger blood meal) [5,23], and have played a principal role in all louse-borne outbreaks investigated through human history, this does not preclude head lice as additional vectors [24]. Moreover, in the last few decades, the status of head lice as a vector of pathogens has been raised, since body louse-borne pathogens have been increasingly detected in head lice collected worldwide, particularly in poor African countries, but also in the USA and France [4,10,12,14,25–27]. This is the case of B. quintana DNA found in head lice belonging to Clade A, E, C and D [6,10,14,25,28,29]. Other pathogens, such as B. recurrentis, Y. pestis and several Acinetobacter species have also been detected in PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 2 / 18 115 / 285 Detection of bacterial pathogens in head lice from Mali human head lice [12,29–31]. Furthermore, experimental infections with R. prowazekii have shown that head lice can be readily infected and disseminate these pathogen in their feces, demonstrating that these lice have the potential to be a vector pathogen under optimal epide- miologic conditions [24]. Emerging infectious diseases represent a challenge for global economies and public health. In remote and underdeveloped regions of the African continent, often no attention is paid towards possible cases of infectious disease until a threshold of serious cases and deaths appears in a cluster and certain epidemic properties are reached. Lice infestations always occurs through blood meals, and the louse remains infected for its entire short life (one month), witnessing a recent human infection [1,32].The usefulness of PCR in detecting bacte- rial DNA in lice has been demonstrated by several investigations. Furthermore, several reports have demonstrated that the study of lice and associated pathogens can be used to detect infected patients (xenodiagnosis), estimate the risk for outbreaks, follow the progress of epi- demics, and justify the implementation of control measures to prevent the spread of infection [32,33]. Our laboratory’s experience in Burundi is the best example of this. Thus, in 1995, D. Raoult et al. identified R. prowazekii in lice collected from Burundi jails, an observation that predicted the huge outbreak of epidemic typhus which erupted in refugee camps in Burundi two years later, in 1997 [1,32,33]. The present work contributes towards this approach, by studying the bacterial pathogens associated with head lice collected in two rural villages in Mali, where a high frequency of head lice infestation had previously been reported [34]. Materials and methods Study area, sampling and ethics statement The study was performed in January 2013 in two rural Malian villages, Done´gue´bougou (12˚ 48’85”N 7˚58’22”W) and Zorocoro (12˚44’75”N 80˚04’50”W), situated in close proximity in the Koulikoro region in a savanna zone. Lice were collected from patients presenting at the health centers in these two villages. All sampled individuals were thoroughly examined for the presence of both head and body lice. All visible head lice were removed from the hair using a fine-toothed comb. In total, 259 head lice samples were isolated from 56 individuals in the vil- lage of Done´gue´bougou and 341 head lice were isolated from 61 individuals in the village of Zorocoro. No body lice were found during the examination. General sanitary and hygienic conditions were poor. All the lice were preserved dry in sterile conditions at room temperature and sent to our laboratory in Marseille (France). This study was approved by the Institutional Ethics Committee of the Faculty of Medicine of Pharmacy and Odontostomatology (permit no. 2013/113/CE/FMPOS). Written informed consent was obtained from the individuals involved or from their legal representatives in the case of children. The representatives of a local health center and the village elders accompanied the researchers for the duration of the study. DNA preparation Prior to DNA isolation and in order to avoid external contamination, the surface of each louse was decontaminated as described previously [18], then each specimen was cut longitudinally into halves. One half was placed in a sterile tube and frozen for later use. The other was crushed in sterile Eppendorf tube and total-DNA was extracted using a DNA extraction kit, QIAamp Tissue Kit (Qiagen, Courtaboeuf, France) using the EZ1 apparatus following the manufacturer’s protocols and stored at 4˚C until use in PCR amplifications. PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 3 / 18 116 / 285 Detection of bacterial pathogens in head lice from Mali Molecular detection of the presence of pathogen DNA Screening of pathogen DNA by qPCR. All DNA samples were screened using quantita- tive real-time PCR (qPCR) using previously reported primers and probes targeting the 16S rRNA gene of Borrelia spp. [35], the 23S gene of Anaplasmataceae spp. [36], the gltA gene of Rickettsia spp.[37], the ompB gene of R. prowazekii [38], the pla gene of Yersinia pestis [38], the yopP gene of B. quintana [25] and the IS1111 of C. burnetii [39]. In addition, all B. quintana and C. burnetii positive samples were confirmed by a second specific qPCR process targeting the fabF3 gene and IS30A spacers respectively [25,39]. All the sequences of primers and probes used for qPCRs and conventional PCRs in this study are given in the supplementary files (S1 Table). All qPCRs were performed using a CFX96™ Real-Time system (Bio-Rad Laboratories, Fos- ter City, CA, USA). The final reaction volume of 20 μl contained 5 μl of the DNA template, 10 μl of Eurogentec™ Probe PCR Master Mix (Eurogentec, Liège, Belgium), 0.5 μM of each primer and 0.5 μM of the FAM-labeled probe. The thermal cycling conditions included one incubation step at 50˚C for two minutes and an initial denaturation step at 95˚C for three min- utes, followed by 40 cycles of denaturation at 95˚C for 15 seconds and annealing extension at 60˚C for 30 seconds. We included the DNA of each target bacteria as positive controls and master mixtures as negative controls to validate each PCR run. No amplifications were detected among the nega- tive controls throughout the study. We considered samples to be positive when the cycle threshold (Ct) was lower than 35 Ct [40]. Conventional PCR and sequencing. All samples that tested positive using Rickettsia genus-specific primers were subjected to standard PCR targeting a 1177-bps fragment of gltA gene [41]. For the identification of Anaplasmataceae species, all positive samples were tested in two PCRs using a set of Anaplasma genus-specific primers targeting the 525-bps fragment of the rpoB gene and a Ehrlichia genus-specific set of primers targeting the 590-bps portion of the groEL gene (heat shock protein gene) [36]. In addition, we determined the multi-spacer typing (MST) of C. burnetii positive samples by amplifying three intergenic spacers (Cox2, Cox5 and Cox18) [42]. All PCR amplification was performed using a Peltier PTC-200 model thermal cycler (MJ Research Inc., Watertown, MA, USA). Reactions were carried out using the Hotstar Taq- polymerase (Qiagen), in accordance with the manufacturer’s instructions. Negative and posi- tive controls were included in each assay. The success of amplification was confirmed by elec- trophoresis on a 1.5% agarose gel. Purification of PCR products was performed using NucleoFast 96 PCR plates (Macherey- Nagel EURL, Hoerdt, France) as per the manufacturer’s instructions. The amplicons were sequenced using the Big Dye Terminator Cycle Sequencing Kit (Perkin Elmer Applied Biosys- tems, Foster City, CA) with an ABI automated sequencer (Applied Biosystems). The electro- pherograms which were obtained were assembled and edited using ChromasPro software (ChromasPro 1.7, Technelysium Pty Ltd., Tewantin, Australia) and compared with those avail- able in the GenBank database by NCBI BLAST (http://blast.ncbi.nlm.nih.gov/Blast.cgi). Mitochondrial clade of lice Determination of louse mitochondrial clade by qPCR assays. To determine the mito- chondrial clades of the lice studied, all the DNA samples were analyzed using clade-specific qPCR assays that targeted a portion of the cytochrome b (cytb) gene, specific to clades A, D, B and C described in our previous study [12]. It is important to note that when we performed the design of the qPCR specific to clade C, clade E was classified as a sub-clade within clade C PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 4 / 18 117 / 285 Detection of bacterial pathogens in head lice from Mali [12], therefore, this qPCR detected both clades C and E. To discriminate between them, we performed another qPCR essay specific only to clade E, targeting 129-bp of cytb (nucleotide position 605–734 of cytb gene). The design was performed and optimized for specificity and sensitivity as described previously[12]. Subsequently, all the lice qPCR which were clade C+E positive were further subjected to qPCR which were clade E specific. We used lice with previ- ously identified clades as positive controls. Negative controls were included in each assay. Cytochrome b amplification and sequencing. For phylogenetic study, forty-five head lice of the total collected were randomly selected and subjected to standard PCR targeting a 347-bp fragment of the cytb gene using the primers and conditions as previously described [8]. Successful amplification was confirmed via gel electrophoresis and amplicons were prepared and sequenced using similar methods as described above for bacteria. Testing of blood meals in head lice For blood meal identification, only head lice specimens with positive bacterial-DNA results were assayed by PCR using the vertebrate-universal specific primers 16SA and 16SB (Table 1) to amplify a 580-bps fragment of the vertebrate host mitochondrial 16S ribosomal RNA as described previously [43]. Successful amplification was confirmed via gel electrophoresis and amplicons were prepared and sequenced using similar methods as described above for bacteria. Data analysis For the head lice cytb sequences obtained in this study, unique haplotypes were defined using DnaSPv5.10 and compared with all the reference cytb haplotypes as described previously [12]. All obtained sequences of Rickettsia and Anaplasmataceae species were analyzed using BLAST (www.ncbi.nlm.nih.gov/blast/Blast.cgi) and compared to sequences in the GenBank database. For C. burnetii, all the sequences obtained from the three spacers were compared with those reported in the reference database available on the website (http://ifr48.timone.univ-mrs.fr/ MST_Coxiella/mst). Sequences of three spacers from all available genotypes were concatenated and aligned using CLUSTAL W for multisequence alignment implemented in MEGA software version 6.06[44]. A maximum-likelihood method was used to infer the phylogenetic analyses and tree recon- struction was performed using MEGA software version 6.06 [44]. Results Lice clade and phylogenetic analysis In total, 600 head lice were collected from 117 individuals living in two villages in Mali and all were tested by qPCRs to determine their clade. Our results show that all the head lice tested (600/600; 100%) belonged to clade E. For phylogenetic study, a total of 45 head lice cytb sequences were analyzed, defining seven different haplotypes, of which four are novel, referred to here E48, E49, E50 and E51, while the remaining three haplotypes possessed the E39 (previously referred as C39) haplotype from Mali and Senegal, and E46 and E47 (previously referred as C46 and C47) haplotypes from Mali. Haplotype E48 was the most prevalent (33.3%), followed by haplotype E39 (26.6%). All the identified haplotypes, together with references from the body and head lice haplogroups were used to construct a maximum-likelihood (ML) tree (Fig 1). All the Malian head lice cytb sequences were clustered with clade E from West Africa (Senegal and Mali). The novel haplo- type sequences identified have been deposited in GenBank (Table 1). PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 5 / 18 118 / 285 Detection of bacterial pathogens in head lice from Mali Table 1. Haplotype frequency of Mali head lice identified per village. Haplotype Done´gue´bougou Zorocoro Total Acc. no. E39 5 1 6 KM579560 E46 3 9 12 KX249780 E47 6 2 8 KX249781 E48 5 10 15 KY937987 E49 1 1 2 KY937988 E50 0 1 1 KY937989 E51 0 1 1 KY937990 Total 20 25 45 https://doi.org/10.1371/journal.pone.0184621.t001 Molecular detection of bacterial pathogens In this study, the qPCR investigation of all 600 head lice samples for Borrelia spp., R. prowaze- kii and Y. pestis produced no positive results. However, we obtained positive results when test- ing for the presence of B. quintana, C. burnetii, Rickettsia spp. and Anaplasmataceae species. Fig 1. Phylogenetic tree showing the relationship between haplotypes identified in this study with other Pediculus humanus haplotypes. The cytb sequences were aligned using CLUSTALW, and phylogenetic inferences were conducted in MEGA 6 using the maximum likelihood method based on the Kimura 2-parameter for nucleotide sequences. The GenBank accession numbers are indicated at the end. Statistical support for the internal branches of the trees was evaluated by bootstrapping with 1,000 iterations. The codon positions included were 1st+2nd+3rd+Noncoding. There was a total of 270 positions in the final dataset. The scale bar represents a 1% nucleotide sequence divergence. https://doi.org/10.1371/journal.pone.0184621.g001 PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 6 / 18 119 / 285 Detection of bacterial pathogens in head lice from Mali The DNA of B. quintana was detected in three of 600 (0.5%) head lice collected from two of 117 (1.7%) persons. All the infected lice were from the village of Done´gue´bougou. No positive samples were found in the village of Zorocoro. Seven of 600 (1.16%) lice samples collected from six of 117 (5.1%) persons tested positive by qPCR using both systems for the presence of C. burnetii DNA. Four of the seven (57.14%) pos- itive lice were from Zorocoro and the remaining three (42.85%) positive lice were from Done´- gue´bougou. We performed MST genotyping of C. burnetii positive lice. Genotype 35, previously recorded in Senegal, was found in one louse from Zorocoro. Another new genotype (genotype 59) was found in two lice from Done´gue´bougou. The phylogenetic position of these genotypes is shown in Fig 2. Interestingly, these two lice were collected from the same person. All attempts to genotype the other positive samples were unsuccessful, possibly because of low DNA concentration. Fig 2. Phylogenetic position of identified genotypes of C. burnetii, the agent of Q fever. The concerned sequences (COX2, 5 and 18) were aligned using CLUSTALW, and phylogenetic inferences was conducted in MEGA 6 using the maximum likelihood method, with the complete deletion option, based on the Kimura 2-parameter for nucleotide sequences. There was a total of 1,247 positions in the final dataset. https://doi.org/10.1371/journal.pone.0184621.g002 PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 7 / 18 120 / 285 Detection of bacterial pathogens in head lice from Mali Table 2. Summary of the pathogens detected in head lice collected from infested individuals in two rural villages in Mali, 2013. Done´gue´bougou Zorocoro Total Bacterial species Persons Head lice Persons Head lice Persons Head lice N = 56 N = 259 N = 61 N = 341 N = 117 N = 600 B. quintana 2 3 0 0 2 (1.7%) 3 (0.5%) C. burnetii 3 3 3 4 6 (5.1%) 7 (1.16%) R. aeschlimannii 1 1 2 3 3 (2.56%) 4 (0.6%) Ehrlichia 6 9 4 5 10 (8.54%) 14 (2.3%) Anaplasma 2 2 0 0 2 (1.7%) 2 (0.3%) Borrelia 0 0 0 0 0 0 Y. pestis 0 0 0 0 0 0 R. prowazekii 0 0 0 0 0 0 https://doi.org/10.1371/journal.pone.0184621.t002 Rickettsial DNA was detected by qPCR targeting the gltA gene in four of 600 (0.6%) head lice collected from three of 117 (2.56%) persons (Table 2). All positive samples were also ampli- fied by conventional PCR using primers targeting a 1,158-bps fragment of the same gene. Three of the four obtained sequences (sample vouchers: Z62HL3, Z62HL4 and D3HL13) were 100% identical to one another, differing by one nucleotide base from the fourth sequence (sample voucher: Z2HL24) and were identified as R. aeschlimannii based on a BLAST search, sharing 99, 65% (1,154 of 1,158 base positions in common) and 99, 74% (1,155 of 1,158 base positions in common) similarity with a reference strain of R. aeschlimannii isolate Crimea-4 (GenBank number KU961540), respectively. These results were also confirmed by a specific qPCR for R. aeschlimannii [39]. The phylogenetic position of this Rickettsia is given in Fig 3. The partial nucleotide sequence of the gltA gene obtained in this study was deposited in the GenBank under accession number: KY937991- KY937992. For Anaplasmataceae, the 23S-based qPCR screening showed 15 out of 600 (2.5%) head lice, collected from 11 of 117 (9.4%) individuals, contained DNA of the Anaplasmataceae spe- cies. Conventional PCR and sequencing using specific Ehrlichia genus-primers targeting a 590-bps fragment of groEL gene showed that 14 of the 15 lice tested were positive for Ehrlichia. Comparison with the GenBank database sequences showed that 11/14 of these sequences form new genotypes closely related to not officially recognized species E. mineirensis UFMG-EV (GenBank number JX629806) with 98.26–98.6% similarities. These new genotypes, referred to here as E. aff. mineirensis, together with E. mineirensis are clustered within the clade of E. canis, as shown in the phylogenetic tree (Fig 4). For three of the 14 remaining sequences, BLAST analysis showed a homology score of under 93% which means that these sequences are likely to be a potential undescribed new species. The closest officially recognized species is E. ewingii (GenBank number AF195273) with 90.4% identity. These three sequences have one to two SNP between them. In the phylogenetic tree (Fig 4), the sequences of this potential new Ehrli- chia sp., provisionally referred to here as “Mali” form a separate and well-supported (bootstrap value 88) branch, which clustered together within the clade that contains E. ewingii from human and other uncultured Ehrlichia sp. from hard ticks. Using the specific Anaplasma genus-primers targeting 525-bps fragment of the rpoB gene, we found that two of the 15 lice tested, collected from two individuals, were positive for Ana- plasma spp. Interestingly, one of the positive lice was also co-infected with E. aff. mineirensis. All the infested lice were from Done´gue´bougou. No positive sampled were found in Zorocoro. A BLAST search showed that these sequences probably belong to an undescribed species, Ana- plasma sp., provisionally referred to here as “Mali”, because only 83% (327/395-bps), 81% (317/392-bps), 80% (316/394-bps) and 80% (315/392-bps) similarities were observed, PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 8 / 18 121 / 285 Detection of bacterial pathogens in head lice from Mali Fig 3. Phylogenetic tree highlighting the position of Rickettsia spp. identified in the present study compared to other Rickettsia bacteria available on GenBank. The gltA sequences were aligned using CLUSTALW, and phylogenetic inferences were conducted in MEGA 6 using the maximum likelihood method, with the complete deletion option, based on the Kimura 3-parameter for nucleotide sequences. The GenBank accession numbers are indicated at the end. Statistical support for the internal branches of the trees was evaluated by bootstrapping with 1,000 iterations. The codon positions included were 1st+2nd+3rd+Noncoding. There was a total of 1,161 positions in the final dataset. The scale bar represents a 2% nucleotide sequence divergence. https://doi.org/10.1371/journal.pone.0184621.g003 respectively, with the rpoB gene of A. phagocytophilum (GenBank number FLME02000004), A. centrale (GenBank number CP001759), A. marginale (GenBank number CP001079) and A. platys (GenBank number KX155493). The phylogenetic position of these Anaplasma are given in Fig 5. Accordingly, E. aff. mineirensis showed an infection rate of 1.8% (11/600) of the total num- ber of lice tested, Ehrlichia sp. “Mali” showed an infection rate of 0.5% (3/600) lice tested and Anaplasma sp. “Mali” showed an infection rate of 0.5% (2/600) lice tested, including one co- infection E. aff. mineirensis/Anaplasma sp. “Mali”. The partial nucleotide sequence of the groEl gene of Ehrlichia and rpoB gene of Anaplasma obtained in this study were deposited in the GenBank under the accession numbers KY937978- KY937986. Blood meal identification in head lice We also performed blood meal analysis in the 29 head lice specimens which were positive for at least one pathogen tested. As expected, DNA from human blood was detected in all lice PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 9 / 18 122 / 285 Detection of bacterial pathogens in head lice from Mali Fig 4. Phylogenetic tree highlighting the position of Ehrlichia spp. identified in the present study compared to other Ehrlichia bacteria available on GenBank. The groEl sequences were aligned using CLUSTALW, and phylogenetic inferences was conducted in MEGA 6 using the maximum likelihood method based on the Kimura 3-parameter model for nucleotide sequences. The GenBank accession numbers are indicated at the end. Statistical support for the internal branches of the trees was evaluated by bootstrapping with 1,000 iterations. The codon positions included were 1st+2nd+3rd+Noncoding. There was a total of 570 positions in the final dataset. The scale bar represents a 5% nucleotide sequence divergence. https://doi.org/10.1371/journal.pone.0184621.g004 tested. Thus, 25 of the 29 obtained sequences showed 100% identity, while the remaining four sequences showed 99.83–99.65% similarities with the 16S ribosomal RNA of Homo sapiens mitochondrial sequences available in the Genbank database. Discussion Human lice infestation remains prevalent worldwide. Surprising and novel insights into the evolution of these ancient and highly intimate scourges of the human race, their bacterial dis- ease agents, and the epidemiology of louse-borne diseases are stimulating a renewal of interest in these bloodsucking insects. Here we provide results of head lice screening from two rural villages of Mali in the savanna zone, where a high rate infestation had previously been reported in 88% of the 112 individuals studied, reflecting the low socioeconomic level in this area [34]. During an epidemiological investigation in New York, the distribution of head lice was associated with gender (boys’ heads are usually shaved), age, socioeconomic status, crowding, methods of storing garments, and family size [5]. Poverty and ignorance appeared to contribute to the persistence of the PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 10 / 18 123 / 285 Detection of bacterial pathogens in head lice from Mali Fig 5. Phylogenetic tree highlighting the position of Anaplasma spp. identified in the present study compared to other Ehrlichia bacteria available on GenBank. The rpoB sequences were aligned using CLUSTALW, and phylogenetic inferences were conducted in MEGA 6 using the maximum likelihood method based on the Kimura 3-parameter for nucleotide sequences. The GenBank accession numbers are indicated at the end. Statistical support for internal branches of the trees was evaluated by bootstrapping with 1,000 iterations. The codon positions included were 1st+2nd+3rd +Noncoding. There was a total of 429 positions in the final dataset. The scale bar represents a 10% nucleotide sequence divergence. https://doi.org/10.1371/journal.pone.0184621.g005 disease [5]. The mtDNA analysis of the 600 head lice collected from 117 Malian individuals, showed that all the head lice tested belonged to clade E, specific to West Africa, as reported by others [6,34]. B. quintana, the causative agent of trench fever, has a long history of association with humans dating back over 4,000 years [45]. Infection was common in France in the 18th cen- tury, during Napoleon’s Russian war, and during World Wars I and II [1,5,46]. It is currently regarded as a re-emerging pathogen in poor countries, as well as in developed countries among the homeless population, where it is responsible for a range of clinical manifestations in humans, including asymptomatic chronic bacteremia, endocarditis and bacillary angioma- tosis [1,5]. It is a very common cause of endocarditis in North Africa [47,48]. For a long time, it was thought that B. quintana was only transmitted by body lice in humans. It was, moreover, found in cats [49] and some human cases have been linked to con- tact with kittens (S2 Table)[50]. Furthermore, in recent years, B. quintana-DNA has fre- quently been detected in head lice collected from impoverished populations such as the homeless or Nepalese children living in slums or on the streets, who are usually infested with both head and body lice [27,51,52], as well as in head lice and head louse nits without concur- rent body lice infestation [14,26,28], highlighting the possible role of head lice as an additional vector in the transmission of B. quintana to humans (S2 Table). In this study, we found B. PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 11 / 18 124 / 285 Detection of bacterial pathogens in head lice from Mali quintana DNA in three of the 600 head lice studied with no evidence of body lice. Our work reinforces findings from previous studies that head lice, as is the case of body lice, may act as vectors of B. quintana. For the first time, the presence of B. quintana in Malian head lice has been shown. All the positive lice were collected from two people living in the same village, Done´gue´bougou. No positive samples were found in Zorocoro. The studies conducted in Mali by Sangare´ et al. failed to detect this bacterium in head lice collected from another Malian village, Diankabou, situated in the Sahelian zone [6,34]. Of all these three villages studied, B. quintana was found in only one village, suggesting a local occurrence of this pathogens. All B. quintana positive head lice were clade E, the unique clade found in the studied era. In recent studies from neigh- boring Senegal, B. quintana was also detected in head lice clade C, which is now recognized as clade E, the same clade found in Mali, as well as in head lice belonging to clade A [14,28]. B. quintana has also been reported in head lice clades C and D from Ethiopia and the Democratic Republic of the Congo, respectively [10,29,53], suggesting that all clades of head lice, except clade B from which no infection has been reported to date, have the potential to serve as vec- tors for B. quintana. Given the scale of head lice infestation around the world, it is of para- mount importance to address their competence as potential disease vectors. In this study, we also assessed our collected lice for the presence of C. burnetii, Rickettsia spp and Anaplasmataceae. These bacteria are usually not associated with human lice, so we used additional tools to confirm that the amplified microorganisms were really associated with human lice. We amplified and sequenced louse cytb from each of positive samples and identi- fied the human blood meal inside each arthropod, so we are sure that we amplified these bacte- ria from engorged human lice. Although all these pathogenic bacteria are not correlated with louse transmission, it is feasible that lice can transmit any agent of chronic bacteremia that is ingested with the blood meal and capable of surviving in the insect’s midgut [1]. Furthermore, lice have been demonstrated to be capable of mechanical transmission for virtually all microor- ganisms tested, including C. burnetii and Rickettsia species [1]. C. burnetii, the causative agent of Q fever, is a worldwide zoonotic disease. The bacterium has a wide host range, including wild and domestic mammals, birds, reptiles, and arthropods, mainly ticks [54]. Infection in humans, usually through aerosol inhalation, can be acute or chronic and the disease exhibits a wide spectrum of clinical manifestations [54,55]. Infections with C. burnetii has been reported throughout the African continent with a high prevalence in Senegal, indicating that Q fever should be considered as a significant public health threat in Africa [55,56]. In Mali, only two serological studies have thus far been performed on humans. The first study was conducted by Tissot-Dupont et al. (1995) and found a seroprevalence of 24% in healthy urban-dwelling people [57]. The second study was conducted by Steinmann et al. (2005) and showed that 40% of 156 mainly adult febrile patients had antibodies against C. burnetii, with 10% of positives having a serological profile suggesting acute infection [58]. Most recently, another study performed on 100 febrile Malian patients (in the village of Dia- nkabou) based on qPCR showed no positive results [56]. One molecular study conducted on both head and body lice from Ethiopia showed no evidence of C. burnetii in all the 98 louse pools tested [53]. The findings from our study showed that 1% of 600 head lice tested infesting 5% of 117 persons studied were C. burnetii DNA positive. Four of the seven positive lice were from Zorocoro and the remaining three positive lice were from Done´gue´bougou. To the best of our knowledge, this is the first molecular evidence of the presence of C. burnetii DNA in head lice infesting individuals from Mali. Although human lice are not known vectors of C. burnetii, it has been shown that, under experimental conditions, it is possible to infect body lice with C. burnetii [59]. There is also a field observation that lice collected in a place where an epidemic of Q fever occurred three months previously are capable of transmitting C. burnetii PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 12 / 18 125 / 285 Detection of bacterial pathogens in head lice from Mali to guinea pigs [20,59]. Our results from Mali, together with data from the literature, suggest that the role of human lice in the epidemiology of Q fever should be investigated further. MST genotyping showed the presence of genotype 35 in one louse from Zorocoro. This genotype was also detected previously in West Africa, in febrile patients and ticks from Senegal [55,56]. Another new genotype (genotype 59) was found in two lice collected from the same person in Done´gue´bougou. A phylogenetic tree based on concatenated sequences (Fig 2) shows that this newly found genotype is mostly related to MST genotypes 6 and 7. As reported in the reference database (http://ifr48.timone.univmrs.fr/mst/coxiella_burnetii/strains.html), genotype 7 was detected in human blood from France and Russia, and genotype 6 was detected in ticks from Senegal and clinical human samples (heart valve and human sera) from France, with no available epidemiological data. Rickettsial species are transmitted by hematophagous arthropods, which contain several agents of human disease [60]. R. prowazekii, a member of the typhus group, is the only known species naturally associated with human lice, in which the body louse is the natural vector and the head louse has been proposed as an additional vector, demonstrated only under laboratory conditions [1,24]. We didn’t find this pathogen in the lice we studied. Although the human louse is not a known vector of rickettsiae species belonging to the spotted fevers group (SFGR), an experimental infection demonstrated that the body louse was able to acquire, maintain, and transmit both R. rickettsii and R. conorii, suggesting that it may play a role, under favorable epidemiologic circumstances, in their transmission to humans [22]. In this study, we demonstrate for the first time the presence of R. aeschlimannii-DNA, another mem- ber of SFG, in 0.6% of the 600 head lice collected from 2.56% of 117 Malian individuals. R. aeschlimannii causing spotted fever was first identified in a patient returning from Morocco [60]. In West Africa, including Mali, this rickettsia was mainly detected in Hya- lomma ticks which appear to be the main vectors and reservoirs [60]. Until now, no human cases of R. aeschlimannii-associated spotted fever has been reported from these countries [60]. Our findings show additional evidence of the presence of the species in Mali being detected in human head lice. Within the Anaplasmataceae family, two significant genera Anaplasma and Ehrlichia, are worldwide tick-borne pathogens that can cause serious illness in a variety of hosts, including humans [61]. These pathogens are not frequently reported in West Africa, with most reports concerning veterinary pathogens in tick vectors [62]. To date, no human cases of anaplasmosis have been reported in Mali. In 1992, a serological survey against E. chaffeensis (the agent of human monocytic ehrlichiosis) in human sera from eight African countries, including Mali, indicated that human ehrlichioses might occur on the continent [63] and a case (diagnosed by serology only) was subsequently reported from Mali [64]. E. chaffeensis has also been identified by PCR in 10% of febrile patients in Central Africa [65], and E. ruminantium-like organisms have been implicated in human infections in South Africa [66]. In the present study, the DNA of Anaplasmataceae was detected in 2.5% of 600 head lice, collected from 9.4% of 117 individu- als. To the best of our knowledge, this is the first evidence of the presence of Anaplasmataceae DNA in human head lice. Ehrlichia was detected in 14 of 600 (2.3%) head lice collected from 10/117 (8.54%) individu- als. Specifically, three of 14 Ehrlichia sequences form a potentially undescribed new species, clustered together within the clade containing E. ewingii from human and other uncultured Ehrlichia sp. from hard ticks. The remaining 11 sequences form new genotype closely related to the not officially recognized species E. mineirensis, a new emerging clade of cattle Ehrlichia pathogens within the E. canis group, the etiologic agent of canine monocytic ehrlichiosis. Recent reports suggested that this species might also be a human pathogen [61]. In 2001, two new ehrlichial genotypes of the E. canis group (which includes E. chaffeensis, E. ruminantium, PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 13 / 18 126 / 285 Detection of bacterial pathogens in head lice from Mali E. muris and E. ewingii) were reported in Rhipicephalus muhsamae from Mali and in Hya- lomma truncatum from Niger [67]. Because in this group, as within each group of ehrlichiae, members share homologous surface antigens and thus cross-react extensively in serologic assays, the authors suggested that these two genotypes may also be organisms responsible for serologic cross-reactions, including in serosurveys and case reports of human ehrlichioses in Africa for currently recognized human pathogenic ehrlichiae [67]. Finally, the DNA of a potential new Anaplasma species was detected in two of 600 (1.58%) head lice collected from in two persons. one of the positive lice was also co-infected with E. aff. mineirensis. Blast analysis of the rpoB gene showed that this Anaplasma sp. was significantly different from all other previously reported Anaplasma species. The closest related species was, with 83% similarities, A. phagocytophilum, the causative agent of human granulocytic anaplas- mosis. This species was recently reported in neighboring Senegal [68]. However, the detection of these potential new species has its limitations, as not all previ- ously described species of Ehrlichia and Anaplasma are already molecularly characterized, so the detection of a ‘new’ genotype may, in fact, be the re-discovery of an old, incompletely char- acterized species [62]. Further studies are required to clarify whether these new genetic vari- ants represent a new species. Furthermore, molecular evidence for the presence of the DNA of these bacteria in head lice cannot distinguish between transient infections, pathogens accidentally acquired from the blood of infected individuals, and those established in a competent vector which can maintain and transmit the pathogen. Nevertheless, all pathogenic species from Anaplasma and Ehrlichia genus are known to be obligatory transmitted by arthropods. Further studies are needed to determine whether the head louse can act as a vector of these bacteria species. Conclusions In conclusion, our finding of several Malian head lice which were positive for B. quintana, C. burnetii, R. aeschlimannii, Anaplasma and Ehrlichia is alarming. Currently, our understanding of the role of human head lice in the epidemiology of these emerging pathogenic bacteria is limited. Since head lice feed only on human blood [5,32], an assumption which is further evi- denced in our study by the identification of human blood meal in all head lice with positive bacterial-DNA, the obtained results imply that the acquired infections are from the blood of patients with ongoing bacteremia. Hence, our study provides a starting point for epidemiologi- cal studies in this area and active survey programs should be encouraged. In Mali, as in the case of several poor African countries, the laboratory capacity to diagnose these infections is often lacking, while many potential emerging pathogens of concern might already be infecting humans but have not yet been detected through disease surveillance. The durability of lice as a sample and the ease with which they can be collected and transported to reference laboratories where suitable molecular biological approaches are available, enhance their potential use as an efficient epidemiological witness for the monitoring and surveillance of emerging pathogens circulating in humans that is critical for the prediction of future disease outbreaks and epidemics at an early stage. Supporting information S1 Table. Oligonucleotide sequences of primers and probes used for real-time PCRs and conventional PCRs in this study. (DOCX) PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 14 / 18 127 / 285 Detection of bacterial pathogens in head lice from Mali S2 Table. Source of B. quintana infection in humans. (RTF) Acknowledgments This work was carried out within the framework of the Me´rieux Doctors Chair of the Acade- mies of Medicine and Sciences (Prof. Ogobara K. Doumbo and Prof. Didier Raoult). Author Contributions Conceptualization: Nadia Amanzougaghene, Florence Fenollar, Abdoul Karim Sangare´, Ogo- bara K. Doumbo, Didier Raoult, Oleg Mediannikov. Data curation: Nadia Amanzougaghene. Formal analysis: Nadia Amanzougaghene, Didier Raoult, Oleg Mediannikov. Funding acquisition: Didier Raoult. Investigation: Abdoul Karim Sangare´, Mahamadou S. Sissoko, Ogobara K. Doumbo, Didier Raoult. Methodology: Nadia Amanzougaghene, Florence Fenollar, Abdoul Karim Sangare´, Mahama- dou S. Sissoko, Ogobara K. Doumbo, Oleg Mediannikov. Resources: Didier Raoult. Supervision: Florence Fenollar, Didier Raoult, Oleg Mediannikov. Validation: Florence Fenollar, Didier Raoult, Oleg Mediannikov. Visualization: Florence Fenollar, Didier Raoult, Oleg Mediannikov. Writing – original draft: Nadia Amanzougaghene. Writing – review & editing: Florence Fenollar, Abdoul Karim Sangare´, Mahamadou S. Sis- soko, Ogobara K. Doumbo, Didier Raoult, Oleg Mediannikov. References 1. Raoult D, Roux V. The body louse as a vector of reemerging human diseases. Clin Infect Dis Off Publ Infect Dis Soc Am. 1999; 29: 888–911. https://doi.org/10.1086/520454 PMID: 10589908 2. Reed DL, Smith VS, Hammond SL, Rogers AR, Clayton DH. Genetic analysis of lice supports direct contact between modern and archaic humans. PLoS Biol. 2004; 2: e340. https://doi.org/10.1371/ journal.pbio.0020340 PMID: 15502871 3. Light JE, Toups MA, Reed DL. What’s in a name: the taxonomic status of human head and body lice. Mol Phylogenet Evol. 2008; 47: 1203–1216. https://doi.org/10.1016/j.ympev.2008.03.014 PMID: 18434207 4. Bonilla DL, Durden LA, Eremeeva ME, Dasch GA. The biology and taxonomy of head and body lice— implications for louse-borne disease prevention. PLoS Pathog. 2013; 9: e1003724. https://doi.org/10. 1371/journal.ppat.1003724 PMID: 24244157 5. Brouqui P. Arthropod-borne diseases associated with political and social disorder. Annu Rev Entomol. 2011; 56: 357–374. https://doi.org/10.1146/annurev-ento-120709-144739 PMID: 20822446 6. Sangare´ AK, Boutellis A, Drali R, Socolovschi C, Barker SC, Diatta G, et al. Detection of Bartonella quintana in African body and head lice. Am J Trop Med Hyg. 2014; 91: 294–301. https://doi.org/10. 4269/ajtmh.13-0707 PMID: 24935950 7. Kittler R, Kayser M, Stoneking M. Molecular evolution of Pediculus humanus and the origin of clothing. Curr Biol CB. 2003; 13: 1414–1417. PMID: 12932325 PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 15 / 18 128 / 285 Detection of bacterial pathogens in head lice from Mali 8. Li W, Ortiz G, Fournier P-E, Gimenez G, Reed DL, Pittendrigh B, et al. Genotyping of human lice sug- gests multiple emergencies of body lice from local head louse populations. PLoS Negl Trop Dis. 2010; 4: e641. https://doi.org/10.1371/journal.pntd.0000641 PMID: 20351779 9. Drali R, Boutellis A, Raoult D, Rolain JM, Brouqui P. Distinguishing body lice from head lice by multiplex real-time PCR analysis of the Phum_PHUM540560 gene. PloS One. 2013; 8: e58088. https://doi.org/ 10.1371/journal.pone.0058088 PMID: 23469145 10. Drali R, Shako J-C, Davoust B, Diatta G, Raoult D. A New Clade of African Body and Head Lice Infected by Bartonella quintana and Yersinia pestis-Democratic Republic of the Congo. Am J Trop Med Hyg. 2015; 93: 990–993. https://doi.org/10.4269/ajtmh.14-0686 PMID: 26392158 11. Ashfaq M, Prosser S, Nasir S, Masood M, Ratnasingham S, Hebert PDN. High diversity and rapid diver- sification in the head louse, Pediculus humanus (Pediculidae: Phthiraptera). Sci Rep. 2015; 5: 14188. https://doi.org/10.1038/srep14188 PMID: 26373806 12. Amanzougaghene N, Akiana J, Mongo Ndombe G, Davoust B, Nsana NS, Parra H-J, et al. Head Lice of Pygmies Reveal the Presence of Relapsing Fever Borreliae in the Republic of Congo. PLoS Negl Trop Dis. 2016; 10: e0005142. https://doi.org/10.1371/journal.pntd.0005142 PMID: 27911894 13. Amanzougaghene N, Mumcuoglu KY, Fenollar F, Alfi S, Yesilyurt G, Raoult D, et al. High Ancient Genetic Diversity of Human Lice, Pediculus humanus, from Israel Reveals New Insights into the Origin of Clade B Lice. PloS One. 2016; 11: e0164659. https://doi.org/10.1371/journal.pone.0164659 PMID: 27741281 14. Boutellis A, Veracx A, Angelakis E, Diatta G, Mediannikov O, Trape J-F, et al. Bartonella quintana in head lice from Se´ne´gal. Vector Borne Zoonotic Dis Larchmt N. 2012; 12: 564–567. https://doi.org/10. 1089/vbz.2011.0845 PMID: 22607067 15. Ayyadurai S, Sebbane F, Raoult D, Drancourt M. Body lice, yersinia pestis orientalis, and black death. Emerg Infect Dis. 2010; 16: 892–893. https://doi.org/10.3201/eid1605.091280 PMID: 20409400 16. Houhamdi L, Lepidi H, Drancourt M, Raoult D. Experimental model to evaluate the human body louse as a vector of plague. J Infect Dis. 2006; 194: 1589–1596. https://doi.org/10.1086/508995 PMID: 17083045 17. Raoult D. A Personal View of How Paleomicrobiology Aids Our Understanding of the Role of Lice in Plague Pandemics. Microbiol Spectr. 2016; 4. https://doi.org/10.1128/microbiolspec.PoH-0001-2014 PMID: 27726806 18. La Scola B, Fournier PE, Brouqui P, Raoult D. Detection and culture of Bartonella quintana, Serratia marcescens, and Acinetobacter spp. from decontaminated human body lice. J Clin Microbiol. 2001; 39: 1707–1709. https://doi.org/10.1128/JCM.39.5.1707-1709.2001 PMID: 11325978 19. Houhamdi L, Raoult D. Experimental infection of human body lice with Acinetobacter baumannii. Am J Trop Med Hyg. 2006; 74: 526–531. PMID: 16606978 20. Giroud P, Jadin J. Infection latente et conservation de “Rickettsia burnetii” chez l’homme, le roˆle du pou. Bull Soc Pathol exotique. 1954; 47: 764–765. 21. Houhamdi L, Fournier P-E, Fang R, Raoult D. An experimental model of human body louse infection with Rickettsia typhi. Ann N Y Acad Sci. 2003; 990: 617–627. PMID: 12860699 22. Houhamdi L, Raoult D. Experimentally infected human body lice (pediculus humanus humanus) as vec- tors of Rickettsia rickettsii and Rickettsia conorii in a rabbit model. Am J Trop Med Hyg. 2006; 74: 521– 525. PMID: 16606977 23. Kim JH, Min JS, Kang JS, Kwon DH, Yoon KS, Strycharz J, et al. Comparison of the humoral and cellu- lar immune responses between body and head lice following bacterial challenge. Insect Biochem Mol Biol. 2011; 41: 332–339. https://doi.org/10.1016/j.ibmb.2011.01.011 PMID: 21296152 24. Robinson D, Leo N, Prociv P, Barker SC. Potential role of head lice, Pediculus humanus capitis, as vec- tors of Rickettsia prowazekii. Parasitol Res. 2003; 90: 209–211. https://doi.org/10.1007/s00436-003- 0842-5 PMID: 12783309 25. Angelakis E, Diatta G, Abdissa A, Trape J-F, Mediannikov O, Richet H, et al. Altitude-dependent Barto- nella quintana genotype C in head lice, Ethiopia. Emerg Infect Dis. 2011; 17: 2357–2359. https://doi. org/10.3201/eid1712.110453 PMID: 22172306 26. Angelakis E, Rolain J-M, Raoult D, Brouqui P. Bartonella quintana in head louse nits. FEMS Immunol Med Microbiol. 2011; 62: 244–246. https://doi.org/10.1111/j.1574-695X.2011.00804.x PMID: 21477003 27. Bonilla DL, Kabeya H, Henn J, Kramer VL, Kosoy MY. Bartonella quintana in body lice and head lice from homeless persons, San Francisco, California, USA. Emerg Infect Dis. 2009; 15: 912–915. https:// doi.org/10.3201/eid1506.090054 PMID: 19523290 28. Diatta G, Mediannikov O, Sokhna C, Bassene H, Socolovschi C, Ratmanov P, et al. Prevalence of Bar- tonella quintana in patients with fever and head lice from rural areas of Sine-Saloum, Senegal. Am J Trop Med Hyg. 2014; 91: 291–293. https://doi.org/10.4269/ajtmh.13-0685 PMID: 24799368 PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 16 / 18 129 / 285 Detection of bacterial pathogens in head lice from Mali 29. Piarroux R, Abedi AA, Shako J-C, Kebela B, Karhemere S, Diatta G, et al. Plague epidemics and lice, Democratic Republic of the Congo. Emerg Infect Dis. 2013; 19: 505–506. https://doi.org/10.3201/ eid1903.121542 PMID: 23750356 30. Boutellis A, Mediannikov O, Bilcha KD, Ali J, Campelo D, Barker SC, et al. Borrelia recurrentis in head lice, Ethiopia. Emerg Infect Dis. 2013; 19: 796–798. https://doi.org/10.3201/eid1905.121480 PMID: 23648147 31. Sunantaraporn S, Sanprasert V, Pengsakul T, Phumee A, Boonserm R, Tawatsin A, et al. Molecular survey of the head louse Pediculus humanus capitis in Thailand and its potential role for transmitting Acinetobacter spp. Parasit Vectors. 2015; 8: 127. https://doi.org/10.1186/s13071-015-0742-4 PMID: 25889008 32. Roux V, Raoult D. Body lice as tools for diagnosis and surveillance of reemerging diseases. J Clin Microbiol. 1999; 37: 596–599. PMID: 9986818 33. Fournier P-E, Ndihokubwayo J-B, Guidran J, Kelly PJ, Raoult D. Human pathogens in body and head lice. Emerg Infect Dis. 2002; 8: 1515–1518. https://doi.org/10.3201/eid0812.020111 PMID: 12498677 34. Sangare´ AK, Doumbo SN, Kone´ AK, Thera MA, Dabo A, Brouqui P, et al. [Lice in Mali: frequency of infestation, genotyping, infection rate and case management]. Med Sante Trop. 2015; 25: 189–193. https://doi.org/10.1684/mst.2015.0442 PMID: 26067516 35. Parola P, Diatta G, Socolovschi C, Mediannikov O, Tall A, Bassene H, et al. Tick-borne relapsing fever borreliosis, rural senegal. Emerg Infect Dis. 2011; 17: 883–885. https://doi.org/10.3201/eid1705. 100573 PMID: 21529402 36. Dahmani M, Davoust B, Rousseau F, Raoult D, Fenollar F, Mediannikov O. Natural Anaplasmataceae infection in Rhipicephalus bursa ticks collected from sheep in the French Basque Country. Ticks Tick- Borne Dis. 2017; 8: 18–24. https://doi.org/10.1016/j.ttbdis.2016.09.009 PMID: 27666778 37. Rolain J-M, Stuhl L, Maurin M, Raoult D. Evaluation of antibiotic susceptibilities of three rickettsial spe- cies including Rickettsia felis by a quantitative PCR DNA assay. Antimicrob Agents Chemother. 2002; 46: 2747–2751. https://doi.org/10.1128/AAC.46.9.2747-2751.2002 PMID: 12183224 38. Nguyen-Hieu T, Aboudharam G, Signoli M, Rigeade C, Drancourt M, Raoult D. Evidence of a louse- borne outbreak involving typhus in Douai, 1710–1712 during the war of Spanish succession. PloS One. 2010; 5: e15405. https://doi.org/10.1371/journal.pone.0015405 PMID: 21060879 39. Mediannikov O, Diatta G, Fenollar F, Sokhna C, Trape J-F, Raoult D. Tick-borne rickettsioses, neglected emerging diseases in rural Senegal. PLoS Negl Trop Dis. 2010; 4. https://doi.org/10.1371/ journal.pntd.0000821 PMID: 20856858 40. Sokhna C, Mediannikov O, Fenollar F, Bassene H, Diatta G, Tall A, et al. Point-of-Care Laboratory of Pathogen Diagnosis in Rural Senegal. PLoS Negl Trop Dis. 2013; 7: e1999. https://doi.org/10.1371/ journal.pntd.0001999 PMID: 23350001 41. Roux V, Rydkina E, Eremeeva M, Raoult D. Citrate synthase gene comparison, a new tool for phyloge- netic analysis, and its application for the rickettsiae. Int J Syst Bacteriol. 1997; 47: 252–261. https://doi. org/10.1099/00207713-47-2-252 PMID: 9103608 42. Glazunova O, Roux V, Freylikman O, Sekeyova Z, Fournous G, Tyczka J, et al. Coxiella burnetii geno- typing. Emerg Infect Dis. 2005; 11: 1211–1217. PMID: 16102309 43. Gatesy J, Amato G, Vrba E, Schaller G, DeSalle R. A Cladistic Analysis of Mitochondrial Ribosomal DNA from the Bovidae. Mol Phylogenet Evol. 1997; 7: 303–319. https://doi.org/10.1006/mpev.1997. 0402 PMID: 9187090 44. Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: Molecular Evolutionary Genetics Anal- ysis version 6.0. Mol Biol Evol. 2013; 30: 2725–2729. https://doi.org/10.1093/molbev/mst197 PMID: 24132122 45. Drancourt M, Tran-Hung L, Courtin J, Lumley de H, Raoult D. Bartonella quintana in a 4000-Year-Old Human Tooth. J Infect Dis. 2005; 191: 607–611. https://doi.org/10.1086/427041 PMID: 15655785 46. Raoult D, Dutour O, Houhamdi L, Jankauskas R, Fournier P-E, Ardagna Y, et al. Evidence for louse- transmitted diseases in soldiers of Napoleon’s Grand Army in Vilnius. J Infect Dis. 2006; 193: 112–120. https://doi.org/10.1086/498534 PMID: 16323139 47. Benslimani A, Fenollar F, Lepidi H, Raoult D. Bacterial Zoonoses and Infective Endocarditis, Algeria. Emerg Infect Dis. 2005; 11: 216–224. https://doi.org/10.3201/eid1102.040668 PMID: 15752438 48. Znazen A, Rolain J-M, Hammami N, Kammoun S, Hammami A, Raoult D. High prevalence of Bartonella quintana endocarditis in Sfax, Tunisia. Am J Trop Med Hyg. 2005; 72: 503–507. PMID: 15891120 49. Drancourt M, Moal V, Brunet P, Dussol B, Berland Y, Raoult D. Bartonella (Rochalimaea) quintana infection in a seronegative hemodialyzed patient. J Clin Microbiol. 1996; 34: 1158–1160. PMID: 8727894 PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 17 / 18 130 / 285 Detection of bacterial pathogens in head lice from Mali 50. Raoult D, Drancourt M, Carta A, Gastaut JA. Bartonella (Rochalimaea) quintana isolation in patient with chronic adenopathy, lymphopenia, and a cat. Lancet Lond Engl. 1994; 343: 977. 51. Sasaki T, Poudel SKS, Isawa H, Hayashi T, Seki N, Tomita T, et al. First molecular evidence of Barto- nella quintana in Pediculus humanus capitis (Phthiraptera: Pediculidae), collected from Nepalese chil- dren. J Med Entomol. 2006; 43: 110–112. PMID: 16506456 52. Koehler JE, Sanchez MA, Garrido CS, Whitfeld MJ, Chen FM, Berger TG, et al. Molecular Epidemiology of Bartonella Infections in Patients with Bacillary Angiomatosis–Peliosis. N Engl J Med. 1997; 337: 1876–1883. https://doi.org/10.1056/NEJM199712253372603 PMID: 9407154 53. Cutler S, Abdissa A, Adamu H, Tolosa T, Gashaw A. Bartonella quintana in Ethiopian lice. Comp Immu- nol Microbiol Infect Dis. 2012; 35: 17–21. https://doi.org/10.1016/j.cimid.2011.09.007 PMID: 22019400 54. Eldin C, Me´lenotte C, Mediannikov O, Ghigo E, Million M, Edouard S, et al. From Q Fever to Coxiella burnetii Infection: a Paradigm Change. Clin Microbiol Rev. 2017; 30: 115–190. https://doi.org/10.1128/ CMR.00045-16 PMID: 27856520 55. Mediannikov O, Fenollar F, Socolovschi C, Diatta G, Bassene H, Molez J-F, et al. Coxiella burnetii in humans and ticks in rural Senegal. PLoS Negl Trop Dis. 2010; 4: e654. https://doi.org/10.1371/journal. pntd.0000654 PMID: 20386603 56. Angelakis E, Mediannikov O, Socolovschi C, Mouffok N, Bassene H, Tall A, et al. Coxiella burnetii- positive PCR in febrile patients in rural and urban Africa. Int J Infect Dis IJID Off Publ Int Soc Infect Dis. 2014; 28: 107–110. https://doi.org/10.1016/j.ijid.2014.05.029 PMID: 25245003 57. Dupont HT, Brouqui P, Faugere B, Raoult D. Prevalence of antibodies to Coxiella burnetti, Rickettsia conorii, and Rickettsia typhi in seven African countries. Clin Infect Dis Off Publ Infect Dis Soc Am. 1995; 21: 1126–1133. 58. Steinmann P, Bonfoh B, Pe´ter O, Schelling E, Traore´ M, Zinsstag J. Seroprevalence of Q-fever in febrile individuals in Mali. Trop Med Int Health TM IH. 2005; 10: 612–617. https://doi.org/10.1111/j.1365-3156. 2005.01420.x PMID: 15941426 59. Babudieri B. Q fever: A zoonosis. Adv Vet Sci. 1959; 5: 82–182. 60. Parola P, Paddock CD, Socolovschi C, Labruna MB, Mediannikov O, Kernif T, et al. Update on tick- borne rickettsioses around the world: a geographic approach. Clin Microbiol Rev. 2013; 26: 657–702. https://doi.org/10.1128/CMR.00032-13 PMID: 24092850 61. Rar V, Golovljova I. Anaplasma, Ehrlichia, and “Candidatus Neoehrlichia” bacteria: pathogenicity, biodi- versity, and molecular genetic characteristics, a review. Infect Genet Evol J Mol Epidemiol Evol Genet Infect Dis. 2011; 11: 1842–1861. https://doi.org/10.1016/j.meegid.2011.09.019 PMID: 21983560 62. Ehounoud CB, Yao KP, Dahmani M, Achi YL, Amanzougaghene N, Kacou N’Douba A, et al. Multiple Pathogens Including Potential New Species in Tick Vectors in Coˆte d’Ivoire. PLoS Negl Trop Dis. 2016; 10: e0004367. https://doi.org/10.1371/journal.pntd.0004367 PMID: 26771308 63. Brouqui P, Le Cam C, Kelly PJ, Laurens R, Tounkara A, Sawadogo S, et al. Serologic evidence for human ehrlichiosis in Africa. Eur J Epidemiol. 1994; 10: 695–698. PMID: 7672049 64. Uhaa IJ, MacLean JD, Greene CR, Fishbein DB. A case of human ehrlichiosis acquired in Mali: clinical and laboratory findings. Am J Trop Med Hyg. 1992; 46: 161–164. PMID: 1539750 65. Ndip LM, Labruna M, Ndip RN, Walker DH, McBride JW. Molecular and clinical evidence of Ehrlichia chaffeensis infection in Cameroonian patients with undifferentiated febrile illness. Ann Trop Med Parasi- tol. 2009; 103: 719–725. https://doi.org/10.1179/000349809X12554106963753 PMID: 20030996 66. Louw M, Allsopp MTEP, Meyer EC. Ehrlichia ruminantium, an emerging human pathogen—a further report. South Afr Med J Suid-Afr Tydskr Vir Geneeskd. 2005; 95: 948, 950. 67. Parola P, Inokuma H, Camicas JL, Brouqui P, Raoult D. Detection and identification of spotted fever group Rickettsiae and Ehrlichiae in African ticks. Emerg Infect Dis. 2001; 7: 1014–1017. https://doi.org/ 10.3201/eid0706.010616 PMID: 11747731 68. Djiba ML, Mediannikov O, Mbengue M, Thiongane Y, Molez J-F, Seck MT, et al. Survey of Anaplasma- taceae bacteria in sheep from Senegal. Trop Anim Health Prod. 2013; 45: 1557–1561. https://doi.org/ 10.1007/s11250-013-0399-y PMID: 23553260 PLOS ONE | https://doi.org/10.1371/journal.pone.0184621 September 20, 2017 18 / 18 131 / 285 S1 Table. Oligonucleotide sequences of primers and probes used for quantitative real- time PCRs and conventional PCRs in this study. Target Name Primers (5’-3’) and probes Source Pediculus humanus F_ GATGTAAATAGAGGGTGGTT Cytochrome b Duplex A- R_ GAAATTCCTGAAAATCAAAC [12] D FAM-CATTCTTGTCTACGTTCATATTTGG-TAMRA VIC-TATTCTTGTCTACGTTCATGTTTGA-TAMRA F_ TTAGAGCGMTTRTTTACCC Duplex B- R_ AYAAACACACAAAAMCTCCT [12] C/E FAM-GAGCTGGATAGTGATAAGGTTTAT-MGB VIC-CTTGCCGTTTATTTTGTTGGGGTTT-TAMRA GGT TGG AAT TGG ATA GTG AT Monoplex This GGG TCC ATA AAG AAA TCC G E study FAM- TAG GAG GCT TTG TGT GTC TAT CCT -TAMRA F_GAGCGACTGTAATTACTAATC Cytb [8] R_CAACAAAATTATCCGGGTCC Rickettsia spp F_GTGAATGAAAGATTACACTATTTAT citrate synthase (gltA) RKNDO3 R_GTATCTTAGCAATCATTCTAATAGC [37] FAM-CTATTATGCTTGCGGCTGTCGGTTC-TAMRA F_ATGACCAATGAAAATAATAAT gltA [40] R_CTTATACTCTCTATGTACA Rickettsia prowazekii F_AATGCTCTTGCAGCTGGTTCT rOmpB gene R_TCGAGTGCTAATATTTTTGAAGCA [37] ompB FAM-CGGTGGTGTTAATGCTGCGTTACAACA- TAMRA Yersinia pestis F_ATG GAG CTT ATA CCG GAA AC plasminogen PLA R_GCG ATA CTG GCC TGC AAG [38] activator gene FAM-TCCCGAAAGGAGTGCGGGTAATAGG-TAMRA Borrelia spp F_AGCCTTTAAAGCTTCGCTTGTAG 16S ribosomal RNA Bor16S R_GCCTCCCGTAGGAGTCTGG [35] FAM-CCGGCCTGAGAGGGTGAACGG-TAMRA Anaplasmataceae F_TGACAGCGTACCTTTTGCAT 23S ribosomal RNA TtAna R_GTAACAGGTTCGGTCCTCCA [36] FAM-GGATTAGACCCGAAACCAAG-TAMRA Anaplasma spp Ana-rpoB F_GCTGTTCCTAGGCTYTCTTACGCGA rpoB gene R_AATCRAGCCAVGAGCCCCTRTAWGG [36] Ehrlichia spp groEL Ehr-groEL F_GTTGAAAARACTGATGGTATGCA [36] gene R_ACACGRTCTTTACGYTCYTTAAC Bartonella quintana F_ TAAACCTCGGGGGAAGCAGA Hypothetical yopP R_ TTTCGTCCTCAACCCCATCA [25] intracellular effector FAM-CGTTGCCGACAAGACGTCCTTG-TAMRA 3-oxoacyl-synthase F_ GCGGCCTTGCTCTTGATGA gene fabF3 R_ GCTACTCTGCGTGCCTTGGA [25] FAM-TGCA GCAGGTGGAGAGAACGTG-TAMRA Coxiella burnetii F_CAAGAAACGTATCGCTGTGGC Spacer IS1111 IS1111 R_CACAGAGCCACCGTATGAATC [39] FAM-CCGAGTTCGAAACAATGAGGGCTG-TAMRA Spacer IS30A F_CGCTGACCTACAGAAATATGTCC R_GGGGTAAGTAAATAATACCTTCTGG IS30A [39] FAM-CATGAAGCGATTTATCAATACGTGTATGC- TAMRA Cox2 Cox2 F_CAACCCTGAATACCCAAGGA [41] 132 / 285 R_GAAGCTTCTGATAGGCGGGA Cox5 CAGGAGCAAGCTTGAATGCG Cox5 F_ [41] R_TGGTATGACAACCCGTCATG Cox18 CGCAGACGAATTAGCCAATC Cox18 F_ [41] R_TTCGATGATCCGATGGCCTT Universal vertebrate 16SA-CGCCTGTTTACCAAAAACAT 16S [42] 16S ribosomal RNA 16SB-CCGGTCTGAACTCAGATCACGT S2 Table. Source of B. quintana infection in humans. Source of infection Status Country References Body lice Confirmed (natural vector) Worldwide [1,5,6] Head lice Suspected France, USA, [6,14,25,27,28,52] Ethiopia, Congo RDC, Senegal, Nepal, Madagascar Kittens or cats Suspected France [48,49] Cat fleas Suspected France [68] 133 / 285 Article 6 : Molecular survey of Head and Body lice, Pediculus humanus, in France Publié dans Vector-Borne and Zoonotic Diseases 2018; 18(5):243-251 134 / 285 VECTOR-BORNE AND ZOONOTIC DISEASES Volume 18, Number 5, 2018 ª Mary Ann Liebert, Inc. DOI: 10.1089/vbz.2017.2206 Molecular Survey of Head and Body Lice, Pediculus humanus, in France Kerdalidec Candy,1,2,* Nadia Amanzougaghene,3,* Arezki Izri,1,2 Sophie Brun,1 Re´my Durand,1 Meriem Louni,4 Didier Raoult,3 Florence Fenollar,4 and Oleg Mediannikov3 Abstract Human lice, Pediculus humanus, are obligate blood-sucking parasites. Phylogenetically, they belong to several mitochondrial clades exhibiting some geographic differences. Currently, the body louse is the only recognized disease vector, with the head louse being proposed as an additional vector. In this article, we study the genetic diversity of head and body lice collected from Bobigny, a town located close to Paris (France), and look for louse-borne pathogens. By amplifying and sequencing the cytb gene, we confirmed the presence of clades A and B in France. Besides, by amplifying and sequencing both cytb and cox1 gene, we reported, for the first time, the presence of clade E, which has thus far only been found in lice from West Africa. DNA from Bartonella quintana was detected in 16.7% of body lice from homeless individuals, but in none of the head lice collected from 47 families. Acinetobacter DNA was detected in 11.5% of head lice belonging to all three clades and 29.1% of body lice. Six species of Acinetobacter were identified, including two potential new ones. Acineto- bacter baumannii was the most prevalent, followed by Candidatus Acinetobacter Bobigny-1, Acinetobacter calcoaceticus, Acinetobacter nosocomialis, Acinetobacter junii, and Candidatus Acinetobacter Bobigny-2. Body lice were found to be infected only with A. baumannii. These findings show for the first time, the presence of clade E head lice in France. This study is also the first to report the presence of DNAs of several species of Acinetobacter in human head lice in France. Keywords: Acinetobacter spp., Bartonella quintana, clade E, France, Pediculus humanus Introduction humans, where the females lay eggs at the base of hair shafts (Light et al. 2008b). They have a global distribution and Downloaded by INIST/CNRS from www.liebertpub.com at 06/20/18. For personal use only. ucking lice (Phthiraptera: Anoplura) are obligate affect individuals of broad economic and social status, Sblood-feeding ectoparasites of placental mammals, particularly school-aged children, regardless of living con- including humans (Chosidow 2000, Light et al. 2008b). Two ditions (Chosidow 2000, Izri et al. 2010, Brouqui 2011). recognized genera parasitize humans: Pthirus and Pediculus. Due to reaction to the bites, they can cause an intense pru- Each genus includes one species affecting humans, Pthirus ritus that may lead to irritation and infection (Veracx and pubis (the crab louse) and Pediculus humanus (Chosidow Raoult 2012). In contrast, body lice live in clothing and 2000, Veracx and Raoult 2012). The latter is of great concern multiply when cold, promiscuity, and lack of hygiene are to public health and includes two ecotypes: head lice and present (Badiaga and Brouqui 2012, Veracx and Raoult body lice. Both ecotypes have the same life cycle and need to 2012). They are often found in jails and in unstable coun- take regular blood meals (approximately five times per day), tries, but are also currently reemerging among homeless on human skin to survive a life span of about 4–12 weeks populations in developed countries (Raoult et al. 1997, (Veracx and Raoult 2012). However, they occupy distinct Brouqui 2011, Badiaga and Brouqui 2012, Veracx and ecological niches. Head lice live in the scalp region of Raoult 2012). 1Department of Parasitology-Mycology, AP-HP, Hoˆpital Avicenne, Bobigny, France. 2UMR ‘‘E´ mergence des Pathologies Virales’’ (EPV: Aix-Marseille University–IRD 190–Inserm 1207–EHESP–IHU Me´diterrane´e Infection), Marseille, France. 3IRD, APHM, MEPHI, IHU-Me´diterrane´e Infection, Aix Marseille University, Marseille, France. 4IRD, APHM, VITROME, IHU-Me´diterrane´e Infection, Aix Marseille University, Marseille, France. *These authors contributed equally to this work. 243 135 / 285 244 CANDY ET AL. Mitochondrial genes (cytb and COI) appear to separate lice Materials and Methods into five divergent mitochondrial clades (A, B, C, D, and E) Ethical clearance exhibiting some geographic differences (Ashfaq et al. 2015, Drali et al. 2015, Amanzougaghene et al. 2016b). Head lice The protocol was reviewed and approved by the Comite´ de encompass all clades, while body lice belong only to clades A Protection des Personnes (institutional review board) of the and D (Ashfaq et al. 2015, Drali et al. 2015, Amanzouga- CPP-Ile-de-France X (2017-02) Ethics Committee. Informed ghene et al. 2016b). The clade A is the most common and is consent was obtained from all patients. found around the world, while clade D is only found in the Republic of Congo and Congo Brazaville (Light et al. 2008a, Study area and lice sampling Ashfaq et al. 2015, Drali et al. 2015, Amanzougaghene et al. 2016a). Clade B is found in America, Europe, Australia, and Between September 2015 and December 2016, head lice North and South Africa, and was most recently reported in were collected from patients attending the Avicenne Hospital Israel, on head lice remains dated *2000 years old (Light in Bobigny. One hundred forty-one patients belonging to 47 et al. 2008a, Amanzougaghene et al. 2016b). Clade C has families were enrolled in this study, and 5 head lice were been found in Ethiopia, the Republic of Congo, and in Asia randomly selected per parasitized family (Supplementary (Amanzougaghene et al. 2016a). Last, clade E consists of Table S1; Supplementary Data are available online at www head lice from West Africa (Senegal and Mali) (Amanzou- .liebertpub.com/vbz). Body lice were obtained from two gaghene et al. 2016b). homeless individuals hospitalized in the same facility. In all, Body lice are potentially more harmful than head lice, 16 and 8 body lice were collected, from patients 1 and 2, because of their role as a vector of at least three pathogenic respectively. After collection, lice were immediately frozen bacteria that have killed millions of people: Rickettsia pro- at -80°C and were then transported to the laboratory of wazekii (the causative agent of epidemic typhus), Bartonella Marseille. In total, 235 head lice and 24 body lice were quintana (trench fever), and Borrelia recurrentis (relapsing processed for molecular study. fever) (Raoult and Roux 1999, Veracx and Raoult 2012). Natural and experimental observations have been made that DNA extraction body lice can also transmit Yersinia pestis, the causative agent of plague, and that they may be the pandemic vectors Before DNA isolation and to avoid external contamina- of this agent (Blanc and Baltazard 1942, Houhamdi et al. tion, the surface of each louse was decontaminated as de- 2006). Some other widespread pathogenic bacteria, such as scribed previously (La Scola et al. 2001). DNA was extracted Serratia marcescens, Acinetobacter baumannii,andAcine- using the QIAamp DNA tissue extraction kit (Qiagen, Hil- tobacter lwoffii, have been detected in human body lice, with den, Germany) in an EZ1 apparatus following the manufac- the assumption that lice may probably also transmit these turer’s instructions. agents to humans (La Scola et al. 2001, Houhamdi and Raoult 2006). Genotypic status of lice Although body lice are currently assumed to be more potent vectors of pathogens, the vector potential of head Identification of louse mitochondrial clade by qPCR as- lice is not yet fully understood. Studies have demonstrated says. To identify the mitochondrial clades of the lice, all that the immune reactions of head lice to different patho- DNA samples were analyzed using clade-specific quantita- gens are stronger than those of body lice, which obviously tive real-time PCR (qPCR) assays that targeted a portion of may carry a broad spectrum of pathogens (Previte et al. cytochrome b (cytb) gene specific to clades A, D, B, and C, as 2014, Kim et al. 2017). In laboratory-reared lice, it has previously described (Amanzougaghene et al. 2016a). It is been demonstrated that head lice can support a persistent important to note that, when the design of the qPCR specific to clade C was performed, clade E was classified as subclade Downloaded by INIST/CNRS from www.liebertpub.com at 06/20/18. For personal use only. load of B. quintana for several days following acquisition in a bloodmeal (Previte et al. 2014). An experimental in- within clade C; therefore, this qPCR detects both clades C fection with R. prowazekii has also shown that head lice and E (Amanzougaghene et al. 2016a). To discriminate be- can be readily infected and disseminate these pathogens in tween them, we performed another qPCR essay, specific only their feces, showing that these lice might be a vector of to clade E (Amanzougaghene et al. 2017). We used lice with pathogens under optimal epidemiologic conditions (Ro- known clades as positive controls and master mixtures as a binson et al. 2003). Indeed, a substantial number of studies negative control for each test. have reported body louse-borne pathogens on head lice collected from different parts of the world. This is the case PCR amplifications and sequencing. For phylogenetic of B. quintana, B. recurrentis,andY. pestis DNA, found in study, 79 (79/235) head lice and 5 (5/24) body lice were head lice belonging to different mitochondrial clades randomly selected and subjected to standard PCR, targeting a (Angelakis et al. 2011a, 2011b, Boutellis et al. 2012, 2013, 347-bp fragment of the cytb gene as previously described (Li Drali et al. 2015, Amanzougaghene et al. 2016a). Several et al. 2010). To confirm the presence of clade E, sixteen lice Acinetobacter species have also been detected in human already identified as clade E by the previous PCR were head lice (Sunantaraporn et al. 2015, Amanzougaghene subjected to another standard PCR, targeting another mito- et al. 2016a). chondrial gene, cytochrome oxidase subunit 1 (cox1), as In this study, we examined the genetic diversity of head previously described (Drali et al. 2016). and body lice collected from Bobigny, a town located 3 km PCR amplification was performed in a Peltier PTC-200 north of Paris (France) (48°54¢38†N2°26¢23†E), to look for model thermal cycler (MJ Research, Inc., Watertown, USA). louse-borne pathogens in these lice. The reactions were carried out using the Hotstar Taq 136 / 285 MOLECULAR SURVEY OF HEAD AND BODY LICE IN FRANCE 245 polymerase (Qiagen), in accordance with the manufacturer’s Cycle Sequencing Kit (Perkin Elmer Applied Biosystems, instructions. Purification of PCR products was performed Foster City, CA) with an ABI automated sequencer (Applied using NucleoFast 96 PCR plates (Macherey-Nagel EURL, Biosystems). The electropherograms were assembled and Hoerdt, France) as per the manufacturer’s instructions. The edited using ChromasPro software (ChromasPro 1.7; Tech- amplicons were sequenced using the Big Dye Terminator nelysium Pty Ltd., Tewantin, Australia). Table 1. Oligonucleotide Sequences of Primers and Probes Used for Real-Time PCRs and Conventional PCRs in This Study Target Name Primers (5¢-3¢) and probes Source Pediculus humanus Duplex F_GATGTAAATAGAGGGTGGTT Amanzougaghene Cytochrome b A–D R_GAAATTCCTGAAAATCAAAC et al. (2016a) FAM-CATTCTTGTCTACGTTCATA TTTGG-TAMRA VIC-TATTCTTGTCTACGTTCATGTTT GA-TAMRA Duplex F_TTAGAGCGMTTRTTTACCC B–C/E R_AYAAACACACAAAAMCTCCT FAM-GAGCTGGATAGTGATAAGGTTT AT-MGB VIC-CTTGCCGTTTATTTTGTTGGGGTT T-TAMRA Monoplex E GGTTGGAATTGGATAGTGAT Amanzougaghene GGGTCCATAAAGAAATCC G et al. (2017) FAM- TAGGAGGCTTTGTGTGTCTATC CT-TAMRA Cytb F_GAGCGACTGTAATTACTAATC Li et al. (2010) R_CAACAAAATTATCCGGGTCC Cytochrome oxidase subunit I Cox1 F_TTAGGGGGTGGTGATCCTGT Drali et al. (2016) R_TGAGAGTGCATTCTTGCTGGT Acinetobacter spp. rpoB F_TACTCATATACCGAAAAGAAACGG Bouvresse et al. (2011) RNA polymerase b R_GGYTTACCAAGRCTATACTCAAC subunit gene FAM-CGCGAAGATATCGGTCTSCAAG C-TAMRA rpoB F_TAYCGYAAAGAYTTGAAAGAAG La Scola et al. (2006) (zone1) R_CMACACCYTTGTTMCCRTGA Rickettsia prowazekii ompB F_AATGCTCTTGCAGCTGGTTCT Nguyen-Hieu et al. (2010) rOmpB gene R_TCGAGTGCTAATATTTTTGAAGCA FAM-CGGTGGTGTTAATGCTGCGTTA CAACA-TAMRA Yersinia pestis PLA F_ATGGAGCTTATACCGGAAAC Nguyen-Hieu et al. (2010) Plasminogen activator gene R_GCGATACTGGCCTGCAAG FAM-TCCCGAAAGGAGTGCGGGTAA Downloaded by INIST/CNRS from www.liebertpub.com at 06/20/18. For personal use only. TAGG-TAMRA Borrelia spp. Bor16S F_AGCCTTTAAAGCTTCGCTTGTAG Parola et al. (2011) 16S ribosomal RNA R_GCCTCCCGTAGGAGTCTGG FAM-CCGGCCTGAGAGGGTGAACG G-TAMRA Bartonella quintana yopP F_TAAACCTCGGGGGAAGCAGA Angelakis et al. (2011a) Hypothetical R_TTTCGTCCTCAACCCCATCA intracellular effector FAM-CGTTGCCGACAAGACGTCCTT G-TAMRA 3-oxoacyl-synthase gene fabF3 F_GCGGCCTTGCTCTTGATGA R_GCTACTCTGCGTGCCTTGGA FAM-TGCAGCAGGTGGAGAGAACG TG-TAMRA Anaplasma spp. TtAna F_TGACAGCGTACCTTTTGCAT Dahmani et al. (2017) 23S ribosomal RNA R_TGGAGGACCGAACCTGTTAC FAM-GGATTAGACCCGAAACCAA G-TAMRA Coxiella burnetii IS1111 F_CAAGAAACGTATCGCTGTGGC Mediannikov et al. (2010) Spacers IS1111 R_CACAGAGCCACCGTATGAATC FAM-CCGAGTTCGAAACAATGAGGG CTG-TAMRA 137 / 285 246 CANDY ET AL. Molecular screening for the presence of pathogen DNA named here as E56–E67 (Table 2). The phylogenetic position The qPCRs were performed to screen all lice samples, of these haplotypes is shown in Figure 1. using previously reported primers and probes, for Acineto- The analysis of -cox1 sequences from 16 head lice be- bacter spp., Borrelia spp., B. quintana, Acinetobacter spp., R. longed to clade E, yielded 6 variable positions defining 4 prowazekii, Y. pestis, Coxiella burnetii, and Anaplasma different haplotypes; all were novel named here as E40–E43 spp. (Mediannikov et al. 2010, Nguyen-Hieu et al. 2010, (Table 3). Phylogenetic tree (Fig. 2) showed that all these Angelakis et al. 2011a, Bouvresse et al. 2011, Parola et al. haplotypes clustered with haplotypes from Mali within clade 2011, Dahmani et al. 2017). All B. quintana-positive samples E, confirming the identity of this clade. were confirmed by a second specific qPCR targeting the fabF3 gene (Angelakis et al. 2011a). All sequences of primers Molecular detection of bacterial pathogens and probes used for qPCRs and conventional PCRs in this study are shown in Table 1. In this study, the qPCR investigation of all lice samples for All qPCRs were performed using a CFX96 Real-Time Rickettsia spp., R. prowazekii, Borrelia spp., Y. pestis, C. system (Bio-Rad, Marnes-la-Coquette, France) and the burnetii, and Anaplasma spp. produced no positive results. Eurogentec Master Mix Probe PCR kit (Eurogentec, Lie`ge, However, we obtained positive results when testing for the Belgium). We included the DNA of the target bacteria as presence of B. quintana and Acinetobacter spp. positive controls and master mixtures as negative control for B. quintana DNA was found in 4 of 24 (16.66%) body lice each test. collected from the same homeless patient. The patient was a To identify the species of Acinetobacter spp., all positive 35-year-old man originally from Pakistan, who was hospi- samples from qPCRs were subjected to standard PCR, tar- talized with an unexplained fever at the time of the sample geting a portion of the rpoB gene as described previously (La collection. No head lice were found to be infected with B. Scola et al. 2006). Amplicons were prepared and sequenced quintana. using similar methods as described for the cytb gene for lice Acinetobacter DNA was detected by qPCR targeting the above. rpoB gene in 27 of 235 head lice (11.5%) collected from 8 of 47 families and in 7 of 8 body lice collected from the Data analysis homeless patient 2, which represented a total of 29.1% (7/24) of all body lice tested. Among these 7 body lice Acinetobacter- For the head and body lice cytb and -cox1 sequences, unique DNA positive, 3 lice were also co-infected with B. quintana. haplotypes were defined using DnaSPv5.10 and compared Conventional PCR and sequencing targeting a 350-bp frag- with the cytb and cox1 haplotypes as described previously ment of the same gene was successful only in 23 of the 34 (Amanzougaghene et al. 2016a, Drali et al. 2016). All se- samples that were positive in qPCR. This may be due to the quences of Acinetobacter species were analyzed using BLAST lower sensitivity of standard PCR compared to qPCR. (www.ncbi.nlm.nih.gov/blast/Blast.cgi) and compared against Based on a BLAST search, seven sequences were identi- sequences in the GenBank database. A maximum-likelihood fied as A. baumannii, two sequences as Acinetobacter noso- method was used to infer the phylogenetic analyses, and tree comialis, one sequence as Acinetobacter junii, and three reconstruction was performed using MEGA software version 6.06 (Tamura et al. 2013). Results Table 2. Haplotype Frequency of Head and Body Lice Identified in Bobigny, France, Based Lice clade and phylogenetic analysis on cytb Gene Of the 141 patients (47 families) infested by head lice, the Head Body Downloaded by INIST/CNRS from www.liebertpub.com at 06/20/18. For personal use only. majority were female (sex ratio M/F = 0.1) aged between 2 Haplotype lice lice Total Acc. no. and 53 years. No body lice were found. In total, 235 head lice collected from the 47 families and 24 A5 14 5 19 KM579542 body lice collected from 2 homeless persons were analyzed A17 5 0 5 KM579553 using qPCRs to determine their clade. The result showed that A60 6 0 6 MF672001 A61 3 0 1 MF672002 82 lice (31.8%, [82/258]) belonged to clade A, 42 (16.3%, A62 1 0 1 MF672003 [42/258]) to clade B, and 134 (51.9%, [134/258]) to clade E. B36 22 0 22 KM579559 All the body lice were clade A, while the head lice belonged B39 4 0 4 MF672004 to all three clades (A, B, and E). E56 9 0 9 MF672005 The analysis of 84 cytb sequences yielded 44 variable E57 3 0 3 MF672006 positions defining 18 different haplotypes. Two haplotypes E58 1 0 1 MF672007 belonged to the worldwide haplotypes, A5 (14 head lice se- E59 7 0 7 MF672008 quences and 5 body lice sequences) and A17 (5 head lice E60 1 0 1 MF672009 sequences), within clade A. Three haplotypes, all from head E61 1 0 1 MF672010 lice, also belonging to clade A, were novel and are named E63 4 0 4 MG759552 E64 2 0 2 MG759553 here as A60–A62. Within clade B, two haplotypes were E65 1 0 1 MG759554 found, one belonged to the B36 haplotype, the most wide- E66 1 0 1 MG759555 spread and prevalent in the B haplogroup, the second was E67 1 0 1 MG759556 novel and is referred to here as B39. The remaining 11 hap- Total 79 5 84 lotypes belonged to clade E, all were novel, and they are 138 / 285 MOLECULAR SURVEY OF HEAD AND BODY LICE IN FRANCE 247 FIG. 1. Phylogenetic tree showing the relationship of haplotypes identified in this study with other Pediculus humanus haplotypes. The cytb sequences were aligned using CLUSTALW and phylogenetic inferences were conducted in MEGA 6 using the maximum likelihood method, based on the Kimura 2-parameter. Statistical support for internal branches of the trees was evaluated by bootstrapping with 500 iterations. There was a total of 270 positions in the final dataset. The scale bar represents a 5% nucleotide sequence divergence. Downloaded by INIST/CNRS from www.liebertpub.com at 06/20/18. For personal use only. sequences as Acinetobacter calcoaceticus, sharing 99–100% to here as Candidatus Acinetobacter Bobigny-1 and Candi- identity with their corresponding reference Acinetobacter datus Acinetobacter Bobigny-2. The most closely related species. species are Acinetobacter johnsonii (GenBank number For 5 of the 23 sequences, BLAST analysis showed a CP010350) for Candidatus Acinetobacter Bobigny-1 with homology score of under 95%, meaning that these sequences 94% similarity (314 of 334 base positions in common), and are likely to correspond to new species, provisionally referred Acinetobacter venetianus (GenBank no. LSVC01000004) for Acinetobacter Bobigny-2 with 91.9% similarity (307 of 334 base positions in common). The phylogenetic position of Table 3. Haplotype Frequency of Head Lice these Acinetobacter are shown in Figure 3. Interestingly, Clade E Identified in Bobigny, France, Candidatus Acinetobacter Bobigny-1 was identified in four Based on Cox1 Gene clade E head lice collected from the same family, while Candidatus Acinetobacter Bobigny-2 was found in one clade Haplotype Head lice Acc. no. B head louse from another family. The remaining 5 of 23 (21.7%) sequences, which were E40 3 MG759563 E41 1 MG759564 also rated, resembled Acinetobacter, but were of poor E42 4 MG759565 quality, which is assumed to be due to co-infection with E43 8 MG759566 several Acinetobacter species. The distribution of these Total 16 species according to lice ecotypes and clades are presented in Table 4. The partial rpoB sequences obtained in this study 139 / 285 248 CANDY ET AL. FIG. 2. Phylogenetic tree showing the relationship of haplotypes identified in this study with other P. humanus haplo- types. The Cox1 sequences were aligned using CLUSTALW and phylogenetic inferences were conducted in MEGA 6 using the maximum likelihood method based on the Kimura 2-parameter. Statistical support for internal branches of the trees was evaluated by bootstrapping with 500 iterations. There was a total of 283 positions in the final dataset. The scale bar represents a 2% nucleotide sequence divergence. were deposited in the GenBank under the accession no.: sampling failed to find this clade among the head lice tested, MF672011-MF672020. possibly due to the limited size and region of louse sampling. B. quintana infection is the most common reemerging louse-borne disease in homeless populations living in poor Discussion hygiene conditions in developed countries, where it is re- Louse infestation remains a social and public health con- sponsible for a range of clinical manifestations in humans, in- cern around the world in the 21st century (Bonilla et al. cluding asymptomatic chronic bacteremia, endocarditis, and Downloaded by INIST/CNRS from www.liebertpub.com at 06/20/18. For personal use only. 2013). In this study, the patients affected by pediculosis ca- bacillary angiomatosis (Raoult and Roux 1999, Brouqui 2011, pitis were mainly female (sex ratio M/F = 0.1). During an Badiaga and Brouqui 2012). In this study, we found B. quintana epidemiological investigation in the same area, 70% (402/ DNA in four of 24 body lice collected from one of the hospi- 574) of infested children were girls and the median age was talized homeless patients, consistent with the standard view that 8.9 years (Bouvresse et al. 2012). body lice are the major natural vector for this pathogen, which The mtDNA analysis of the 235 head and 24 body lice is among the most prevalent parasitic infestations in the showed the presence of three major haplogroups: A, B, and E. homeless population (Raoult and Roux 1999, Badiaga et al. Haplogroup E was the most prevalent (51.9%, [134/258]), 2008). Epidemiologic studies of homeless populations have followed by haplogroups A (31.8%, [82/258]) and B (16.3%, reported the prevalence of 7–22% for body louse infestations [42/258]). All the body lice belonged to clade A. These data and 2–30% for B. quintana infections (Badiaga et al. 2008). confirm the existence of clade A and B in France, as reported In recent decades, B. quintana DNA has also been detected by others (Light et al. 2008a, Drali et al. 2015, Amanzouga- in head lice collected from poor and homeless persons in the ghene et al. 2016b). Previous studies reported that clade E was United States, Nepal, Senegal, Ethiopia, the Democratic Re- limited to West African countries, namely Senegal and Mali public of the Congo, and France (Sasaki et al. 2006, Bonilla (Ashfaq et al. 2015, Amanzougaghene et al. 2016b, 2017). et al. 2009, Angelakis et al. 2011a, 2011b, Boutellis et al. 2012, This is the first report of clade E found in head lice in France, Cutler et al. 2012, Drali et al. 2014, 2015). Conversely, all which may be explained by the intercontinental migration attempts to detect B. quintana in head lice collected from flow. Furthermore, clade C lice that were previously only schoolchildren living in family households have failed observed in Africa and Asia have also recently been found in (Fournier et al. 2002, Bouvresse et al. 2011, Sunantaraporn head lice collected in Paris (Drali et al., unpublished data). Our et al. 2015). Taken together, these results suggest that head lice 140 / 285 MOLECULAR SURVEY OF HEAD AND BODY LICE IN FRANCE 249 FIG. 3. Phylogenetic tree highlighting the position of Acinetobacter species identified in head and body lice compared to another Acinetobacter available in the GenBank database. The rpoB sequences were aligned using CLUSTALW, and phylogenetic inferences were conducted in MEGA 6 using the maximum likelihood method based on the Kimura 3- parameter model for nucleotide sequences. The GenBank accession numbers are indicated at the end. Statistical support for internal branches of the trees was evaluated by bootstrapping with 1000 iterations. There was a total of 345 positions in the final dataset. The scale bar represents a 10% nucleotide sequence divergence. probably can also transmit B. quintana if people live close together in poor sanitary conditions and if they lack medical Table 4. Summary of the Bacterial Species treatment. This view is supported by our finding, as we were Detected in Head and Body Lice Collected unable to detect the bacterium in any of the head lice collected from Infested Individuals in Bobigny Per from the 47 middle-class suburban families in the study. Downloaded by INIST/CNRS from www.liebertpub.com at 06/20/18. For personal use only. Lice Ecotype and Clade In this study, we also assessed our collected lice for the presence of Acinetobacter species. More attention is now paid to Head or body Clade of Total extrahospital reservoirs of these ubiquitous opportunistic bac- Bacterial species lice H/B (no.) lice (no.) (%) teria and their potential involvement in emerging human Bartonella quintana B A 4 (1.5) community-acquired infections, as pan drug-resistant strains are increasingly being identified worldwide (Eveillard et al. 2013). Acinetobacter B (4) and A (6) and 7 (2.7) baumannii H (3) B (1) Recent works have shown the Acinetobacter infection to be highly prevalent among body and head lice (La Scola and Acinetobacter H B (1) and 2 (0.7) nosocomialis A (1) Raoult 2004, Bouvresse et al. 2011, Amanzougaghene et al. 2016a). However, it is still unknown how lice acquire these Acinetobacter junii H B 1 (0.4) infections. Some authors argue that infections could occur Acinetobacter H B (1) and 3 (1.15) after the ingestion of an infected blood meal from patients calcoaceticus 2 (E) with ongoing bacteremia (La Scola and Raoult 2004) or Candidatus H E 4 (1.5) possibly by passage through the human skin while feeding Acinetobacter bobigny-1 (Kempf et al. 2012). Furthermore, an experimental study showed that the human body louse, feeding on bacteremic Candidatus H B 1 (0.4) Acinetobacter rabbits, can acquire and support a persistent life-long infec- bobigny-2 tion with A. baumannii and A. lwoffii (Houhamdi and Raoult Acinetobacter spp. H A, B, and E 5 (1.9) 2006). Another study compared two sequenced genomes of A. baumannii and showed that the A. baumannii homeless strain 141 / 285 250 CANDY ET AL. from the body louse had several hundred insertion sequence Authors’ Contributions elements, which played a crucial role in its genome reduction Conceived and designed the experiments: O.M., F.F., and compared to the human multidrug-resistant A. baumannii A.I.. Collected samples: K.C., A.I., and S.B. Conducted the AYE strain, and which have also been shown to have low experiments: K.C., N.A., O.M., and F.F. Analyzed the data: catabolic capacities, suggesting the specific adaptation of this K.C., N.A., and O.M., and F.F. Wrote the article: N.A. K.C., strain to the louse environment (Vallenet et al. 2008). O.M., F.F. A.I., R.D., M.L., and D.R. Our results showed the presence of Acinetobacter DNA in 11.5% of head lice and 29.1% of body lice. Six species were Author Disclosure Statement identified, including two potential new ones. A. baumannii was the most prevalent, followed by Candidatus Acinetobacter No competing financial interests exist. Bobigny-1, A. calcoaceticus, A. nosocomialis, A. junii,and Candidatus Acinetobacter Bobigny-2. Body lice were found to References be infected only by A. baumannii, while the head lice belonging Amanzougaghene N, Akiana J, Mongo Ndombe G, Davoust B, to all three clades were found to be infected by at last one of all et al. Head lice of pygmies reveal the presence of relapsing six species. As a result, it appears that Acinetobacter lice in- fever Borreliae in the Republic of Congo. PLoS Negl Trop fection is not specific to a particular louse clade, but probably to Dis 2016a; 10:e0005142. louse ecotypes, as body lice were found to be infected only by A. Amanzougaghene N, Fenollar F, Sangare´ AK, Sissoko MS, baumannii. This hypothesis is supported by the detection of A. et al. Detection of bacterial pathogens including potential new baumannii in 21% of body lice collected worldwide (La Scola species in human head lice from Mali. PLoS One 2017; 12: and Raoult 2004). In contrast, studies performed on head lice e0184621. collected from elementary school children in Thailand and from Amanzougaghene N, Mumcuoglu KY, Fenollar F, Alfi S, et al. pygmy population in the Republic of Congo, showed the High ancient genetic diversity of human lice, pediculus hu- presence of the DNA of several Acinetobacter species, in 3.62% manus, from Israel Reveals New Insights into the Origin of of head lice belonging to both clade A and C, and 37.3% of head Clade B Lice. PLoS One 2016b; 11:e0164659. lice belonging to clades A, D, and C (Sunantaraporn et al. 2015, Angelakis E, Diatta G, Abdissa A, Trape J-F, et al. Altitude- Amanzougaghene et al. 2016a), respectively. In these two dependent Bartonella quintana genotype C in head lice, studies, the Acinetobacter species identified were A. junii, A. Ethiopia. Emerg Infect Dis 2011a; 17:2357–2359. ursingii, A. baumannii, A. johnsonii, A. schandleri, A. lwoffii, A. Angelakis E, Rolain J-M, Raoult D, Brouqui P. Bartonella nosocomialis, A. towneri,andA. radioresistens (Sunantaraporn quintana in head louse nits. FEMS Immunol Med Microbiol et al. 2015, Amanzougaghene et al. 2016a). When comparing 2011b; 62:244–246. Ashfaq M, Prosser S, Nasir S, Masood M, et al. High diversity the panel of Acinetobacter species found in all these studies with and rapid diversification in the head louse, Pediculus huma- our findings, A. ursingii, A. johnsonii, A. schandleri, A. lwoffii, nus (Pediculidae: Phthiraptera). Sci Rep 2015; 5:14188. A. towneri,andA. radioresistens were not identified in our head Badiaga S, Brouqui P. Human louse-transmitted infectious lice collection. Conversely, our sampling showed, for the first diseases. Clin Microbiol Infect Off Publ Eur Soc Clin Mi- time, the presence of the DNA of A. calcoaceticus and two crobiol Infect Dis 2012; 18:332–337. potentially new species in human head lice. Badiaga S, Raoult D, Brouqui P. Preventing and controlling These studies together demonstrate the widespread infec- emerging and reemerging transmissible diseases in the tion of human lice with several species of Acinetobacter, homeless. Emerg Infect Dis 2008; 14:1353–1359. suggesting that lice could be a preferential host for these Blanc G, Baltazard M. Role of human ectoparasites in the bacteria. However, it remains to be determined whether these transmission of plague. Bull Acad Med 1942; 1126:448. Acinetobacter strains present in lice are the same as those that Bonilla DL, Durden LA, Eremeeva ME, Dasch GA. The biology are responsible for human infections. and taxonomy of head and body lice—implications for louse- Downloaded by INIST/CNRS from www.liebertpub.com at 06/20/18. For personal use only. In conclusion, we confirmed that head lice from Bobigny borne disease prevention. PLoS Pathog 2013; 9:e1003724. (France) belong to haplogroups A and B, and reported, for the Bonilla DL, Kabeya H, Henn J, Kramer VL, et al. Bartonella first time, the presence of haplogroup E, which is specific to quintana in body lice and head lice from homeless persons, West Africa, reflecting the heterogeneous communities San Francisco, California, USA. Emerg Infect Dis 2009; 15: found in the studied area. We detected B. quintana only in 912–915. body lice from homeless individuals, but not from head lice Boutellis A, Mediannikov O, Bilcha KD, Ali J, et al. Borrelia collected from the 47 middle-class suburban families. Sev- recurrentis in head lice, Ethiopia. Emerg Infect Dis 2013; 19: eral Acinetobacter species were also detected, including two 796–798. potentially new ones, indicating that lice could be a source for Boutellis A, Veracx A, Angelakis E, Diatta G, et al. Bartonella quintana in head lice from Se´ne´gal. Vector Borne Zoonotic Acinetobacter spp. infections in humans. Dis Larchmt N 2012; 12:564–567. Bouvresse S, Berdjane Z, Durand R, Bouscaillou J, et al. Per- Acknowledgments methrin and malathion resistance in head lice: Results of ex We gratefully thank IHU Fondation Me´diterrane´e Infec- vivo and molecular assays. J Am Acad Dermatol 2012; 67: tion for supporting the study. 1143–1150. Bouvresse S, Socolovshi C, Berdjane Z, Durand R, et al. No Funding Source evidence of Bartonella quintana but detection of Acinetobacter baumannii in head lice from elementary schoolchildren in This study was supported by Me´diterrane´e Infection and Paris. Comp Immunol Microbiol Infect Dis 2011; 34:475–477. the National Research Agency under the program « In- Brouqui P. Arthropod-borne diseases associated with political vestissements d’avenir », reference ANR-10-IAHU-03. and social disorder. Annu Rev Entomol 2011; 56:357–374. 142 / 285 MOLECULAR SURVEY OF HEAD AND BODY LICE IN FRANCE 251 Chosidow O. Scabies and pediculosis. Lancet Lond Engl 2000; Light JE, Allen JM, Long LM, Carter TE, et al. Geographic 355:819–826. distributions and origins of human head lice (Pediculus hu- Cutler S, Abdissa A, Adamu H, Tolosa T, et al. Bartonella manus capitis) based on mitochondrial data. J Parasitol quintana in Ethiopian lice. Comp Immunol Microbiol Infect 2008a; 94:1275–1281. Dis 2012; 35:17–21. Light JE, Toups MA, Reed DL. What’s in a name: The taxo- Dahmani M, Davoust B, Rousseau F, Raoult D, et al. Natural nomic status of human head and body lice. Mol Phylogenet Anaplasmataceae infection in Rhipicephalus bursa ticks col- Evol 2008b; 47:1203–1216. lected from sheep in the French Basque Country. Ticks Tick- Mediannikov O, Diatta G, Fenollar F, Sokhna C, et al. Tick- Borne Dis 2017; 8:18–24. borne rickettsioses, neglected emerging diseases in rural Se- Drali R, Abi-Rached L, Boutellis A, Djossou F, et al. Host negal. PLoS Negl Trop Dis 2010; 4:e654. switching of human lice to new world monkeys in South Nguyen-Hieu T, Aboudharam G, Signoli M, Rigeade C, et al. America. Infect Genet Evol J Mol Epidemiol Evol Genet Evidence of a louse-borne outbreak involving typhus in Infect Dis 2016; 39:225–231. Douai, 1710–1712 during the war of Spanish succession. Drali R, Sangare´ AK, Boutellis A, Angelakis E, et al. Barto- PLoS One 2010; 5:e15405. nella quintana in body lice from scalp hair of homeless Parola P, Diatta G, Socolovschi C, Mediannikov O, et al. Tick- persons, France. Emerg Infect Dis 2014; 20:907–908. borne relapsing fever borreliosis, rural senegal. Emerg Infect Drali R, Shako J-C, Davoust B, Diatta G, et al. A new clade of Dis 2011; 17:883–885. African body and head lice infected by Bartonella quintana Previte D, Olds BP, Yoon K, Sun W, et al. Differential gene and Yersinia pestis-Democratic Republic of the Congo. Am J expression in laboratory strains of human head and body lice Trop Med Hyg 2015; 93:990–993. when challenged with Bartonella quintana, a pathogenic Eveillard M, Kempf M, Belmonte O, Pailhorie`s H, et al. Re- bacterium. Insect Mol Biol 2014; 23:244–254. servoirs of acinetobacter baumannii outside the hospital and Raoult D, Roux V. The body louse as a vector of reemerging potential involvement in emerging human community- human diseases. Clin Infect Dis Off Publ Infect Dis Soc Am acquired infections. Int J Infect Dis IJID Off Publ Int Soc 1999; 29:888–911. Infect Dis 2013; 17:e802–e805. Raoult D, Roux V, Ndihokubwayo JB, Bise G, et al. Jail fever Fournier P-E, Ndihokubwayo J-B, Guidran J, Kelly PJ, et al. (epidemic typhus) outbreak in Burundi. Emerg Infect Dis Human pathogens in body and head lice. Emerg Infect Dis 1997; 3:357–360. 2002; 8:1515–1518. Robinson D, Leo N, Prociv P, Barker SC. Potential role of head Houhamdi L, Lepidi H, Drancourt M, Raoult D. Experimental lice, Pediculus humanus capitis, as vectors of Rickettsia model to evaluate the human body louse as a vector of plague. prowazekii. Parasitol Res 2003; 90:209–211. J Infect Dis 2006; 194:1589–1596. Sasaki T, Poudel SKS, Isawa H, Hayashi T, et al. First mo- Houhamdi L, Raoult D. Experimental infection of human body lecular evidence of Bartonella quintana in Pediculus huma- lice with Acinetobacter baumannii. Am J Trop Med Hyg nus capitis (Phthiraptera: Pediculidae), collected from 2006; 74:526–531. Nepalese children. J Med Entomol 2006; 43:110–112. Izri A, Uzzan B, Maigret M, Gordon MS, et al. Clinical efficacy Sunantaraporn S, Sanprasert V, Pengsakul T, Phumee A, et al. and safety in head lice infection by Pediculus humanis capitis Molecular survey of the head louse Pediculus humanus ca- De Geer (Anoplura: Pediculidae) of a capillary spray con- pitis in Thailand and its potential role for transmitting Aci- taining a silicon-oil complex. Parasite 2010; 17:329–335. netobacter spp. Parasit Vectors 2015; 8:127. Kempf M, Abdissa A, Diatta G, Trape J-F, et al. Detection of Tamura K, Stecher G, Peterson D, Filipski A, et al. MEGA6: acinetobacter baumannii in human head and body lice from molecular evolutionary genetics analysis version 6.0. Mol Ethiopia and identification of new genotypes. Int J Infect Dis Biol Evol 2013; 30:2725–2729. 2012; 16:e680–e683. Vallenet D, Nordmann P, Barbe V, Poirel L, et al. Comparative Kim JH, Previte DJ, Yoon KS, Murenzi E, et al. Comparison of analysis of acinetobacters: three genomes for three lifestyles. the proliferation and excretion of Bartonella quintana be- PLoS One 2008; 3:e1805. Downloaded by INIST/CNRS from www.liebertpub.com at 06/20/18. For personal use only. tween body and head lice following oral challenge. Insect Veracx A, Raoult D. Biology and genetics of human head and Mol Biol 2017; 26:266–276. body lice. Trends Parasitol 2012; 28:563–571. La Scola B, Fournier P-E, Brouqui P, Raoult D. Detection and culture of Bartonella quintana, Serratia marcescens, and Acinetobacter spp. from decontaminated human body lice. J Address correspondence to: Clin Microbiol 2001; 39:1707–1709. Oleg Mediannikov La Scola B, Gundi VAKB, Khamis A, Raoult D. Sequencing of IRD, APHM, MEPHI, URMITE, the rpoB gene and flanking spacers for molecular identification IHU-Me´diterrane´e Infection of Acinetobacter species. J Clin Microbiol 2006; 44:827–832. Aix Marseille University La Scola B, Raoult D. Acinetobacter baumannii in human body 19-21 Boulevard Jean Moulin louse. Emerg Infect Dis 2004; 10:1671–1673. Marseille 13005 Li W, Ortiz G, Fournier P-E, Gimenez G, et al. Genotyping of France human lice suggests multiple emergencies of body lice from local head louse populations. PLoS Negl Trop Dis 2010; 4:e641. E-mail: [email protected] 143 / 285 Article 7 : Detection of bacterial pathogens in clade E head lice collected from Niger's refugees in Algeria. Publié dans Parasite and Vectors (2018) 11:348 144 / 285 Louni et al. Parasites & Vectors (2018) 11:348 https://doi.org/10.1186/s13071-018-2930-5 RESEARCH Open Access Detection of bacterial pathogens in clade E head lice collected from Niger’s refugees in Algeria Meriem Louni1,2, Nadia Amanzougaghene3, Nassima Mana4, Florence Fenollar2, Didier Raoult3, Idir Bitam2,4,5 and Oleg Mediannikov3* Abstract Background: Head lice, Pediculus humanus capitis, are obligate blood-sucking parasites. Phylogenetically, they occur in five divergent mitochondrial clades (A, D, B, C and E), each having a particular geographical distribution. Recent studies have revealed that head lice, as is the case of body lice, can act as a vector for louse-borne diseases. Here, we aimed to study the genetic diversity of head lice collected from Niger’s refugees (migrant population) arriving in Algeria, northern Africa, and to look for louse-borne pathogens. Comparative head lice samples collected from indigenous population of schoolchildren (non-immigrant) were also analyzed to frame the study. Results: In this study, 37 head lice samples were collected from 31 Nigerien refugees, as well as 45 head lice from 27 schoolchildren. The collection was established in three localities of eastern Algiers, north Algeria. Quantitative real-time PCR screening of pathogens bacteria and the genetic characterisation of the head lice satut were performed. Through amplification and sequencing of the cytb gene, results showed that all head lice of Nigerien refugees 37/82 (45.12%) belonged to clade E with the presence of four new haplotypes, while, of the 45 head lice of schoolchildren, 34/82 lice (41.46%) belonged to clade A and 11/82 (13.41%) belonged to clade B. Our study is the first to report the existence of clade E haplogroup in Nigerien head lice. DNA of Coxiella burnetii was detected in 3/37 (8.10%) of the head lice collected from 3 of the 31 (9.67%) migrant population. We also revealed the presence of Acinetobacter DNA in 20/37 (54.05%) of head lice collected from 25/31 (80.64%) of the Nigerien refugees, and in 25/45 (55.55%) head lice collected from 15/27 (55.55%) schoolchildren. All positive Nigerien-head lice for Acinetobacter spp. were identified as A. baumannii, while positive schoolchildren-head lice were identified as A. johnsonii 15/25 (60%), A. variabilis 8/25 (32%) and A. baumannii 2/25 (8%). Conclusions: Based on these findings from head lice collected on migrant and non-migrant population, our results show, for the first time, that head lice from Niger belong to haplogroup E, and confirm that the clade E had a west African distribution. We also detected, for the first time, the presence of C. burnetii and A. baumannii in these Nigerien head lice. Nevertheless, further studies are needed to determine whether the head lice can transmit these pathogenic bacteria from one person to another. Keywords: Pediculus humanus capitis, Head lice, Niger’s refugees, Scholchildren, Migrant population, Non-migrant population, Coxiella burnetii, Acinetobacter spp., Algeria * Correspondence: [email protected] 3IRD, AP-HM, MEPHI, IHU-Méditerranée Infection, Aix-Marseille Univ, Marseille, France Full list of author information is available at the end of the article © The Author(s). 2018 Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated. 145 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 2 of 11 Background probably, transmitting the causative agent of plague, Yersi- Sucking lice (Phthiraptera: Anoplura) are permanent nia pestis, during plague pandemics [18–20]. Although parasitic insects that infest mammals, including humans body lice are currently presumed to be harmful vectors of [1, 2]. Two recognized genera parasitize humans, Pthirus pathogens and to play a major role in all lice epidemics and Pediculus, both include one species in humans, studied throughout human history, the status of head lice Pthirus pubis and Pediculus humanus [2, 3]. The latter as a vector of louse-borne diseases is still under debate. In- comprises two ecotypes, the head louse (Pediculus deed, several studies report findings of body louse-borne humanus capitis) and the body louse (Pediculus huma- pathogens in head lice belonging to different mitochon- nus humanus). They are obligate blood-feeding parasites drial clades in different parts of the world [11, 12, 16, 21– that thrive exclusively on human blood [4, 5]. The two 25]. Several Acinetobacter species have also been detected ecotypes have the same life-cycle; however, they occupy in human head lice [11, 14, 16, 26, 27]. distinct ecological niches and have different feeding pat- The emergence of humanitarian crises around the terns: head lice live and multiply exclusively on the world, involving thousands of migrants, can represent an scalp, while body lice live and multiply in the clothes explosive risk factor in arthropod-borne disease epi- and feed on the human body [3, 4]. Contrary to body demics. Indeed, several reports of louse-borne B. recur- lice, that are mostly prevalent in people living in precar- rentis have been from various European countries in ious conditions where cold and lack of hygiene are refugee communities migrating from the Horn of Africa present, head lice are prevalent worldwide and preferen- [28–32]. Thus, refugee populations generate a risk of the tially infest schoolchildren, regardless of social class or propagation of lice and their associated bacterial diseases level of hygiene. They can cause very intense pruritus [30]. Algeria continues to receive thousands of people that may lead to high irritation and even wound infec- from different countries, particularly from west Africa tions [6–8]. The head louse is most likely to have been and mostly from Niger. associated with humans since our pre-hominid ancestors In this study, we aimed to assess the occurrence of and has been dispersed throughout the world by human bacterial pathogens associated with head lice collected migration [2]. from Niger’s refugees arriving in Algeria, using molecu- Molecular analysis of the mitochondrial genes cyto- lar tools. The determination of the genetic status of the chrome b (cytb)andcytochrome c oxidase subunit 1 head lice was also performed. Besides, a comparative (cox1) allowed to infer a robust phylogeographical classi- sample of head lice collected from local population fication of P. humanus into five mitochondrial clades (A, (non-migrant) was submitted to the same examination D, B, C and E), each with a particular geographical dis- in the order to frame the results obtained within the tribution [9, 10]. Clade A and D include both head and head lice of Nigerien refugees (migrant population). body lice, in contrast to the other clades that include only head lice, therefore the head lice encompass all the Methods diversity [9, 10]. Clade A is the most common and has a Louse sampling worldwide distribution, while clade D is only found in In January 2016, an epidemiological investigation was central Africa (Ethiopia and the Democratic Republic of established in the Bab Ezzouar Nigerien refugees camp the Congo) [11, 12]. Clade B is confined to America, (36°43′00″N, 3°11′00″E), East Algiers, Algeria. Most refu- Europe, Australia, north and south Africa and was most gees came from Zinder, southern Niger, after stopping in recently reported in head lice remains from Israel, dating Tamanrasset, located in the extreme south of Algeria from approximately 2000 years ago [9, 13]. Clade C is (Fig. 1). All of the sampled individuals were examined for found in Ethiopia, the Democratic Republic of Congo the presence of both head and body lice; however, no body and in Asia (Nepal and Thailand) [11, 13–15]. Lastly, lice were found during the examination. A total of 37 head clade E, a novel clade described in west Africa (Senegal lice were collected from 31 individuals. and Mali) was previously described as a sub-clade of Between November 2015 and May 2016, head lice clade C [9, 10]. Recently, this latter was also reported, were collected from schoolchildren in 5 elementary for the first time, in head lice from Bobigny, France [16]. schools at 3 different location in eastern Algiers, north Until recently, human body lice remained the only Algeria, including the regions of Bab Ezzouar, El main vector of three life-threatening infectious diseases Mohammadia (36°44′00″N, 3°08′00″E,) and Bordj el that have killed millions of people throughout the his- kiffan (36°45′00″N, 3°11′00″E), in order to have a com- tory of humanity namely: epidemic typhus, trench fever parative sample from the local population. A total of 45 and relapsing fever, caused by Rickettsia prowazekii, head lice were collected from 27 schoolchildren (Fig. 2). Bartonella quintana and Borrelia recurrentis, respect- Visible lice were removed from scalps using clamps ively [17]. Natural and experimental observations show and were immediately frozen at -20 °C and then trans- that body lice may also play a role in hosting and, ported to the IHU Méditerrannée-Infection, Marseille. 146 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 3 of 11 Fig. 1 a Refugee camp showing squalor and unhygienic conditions. b Travel routes of Niger’s refugees from Zinder, Niger, western Africa to Algiers, Algeria, northern Africa All head lice collected were then processed for molecu- 95 °C for 15 min, followed by 40 cycles consisting of 1 lar study. min denaturation at 95 °C, 30 s annealing at 55 °C and a 1 min extension at 72 °C. A final extension cycle at 72 °C DNA extraction for 7 min was performed and the reactions were cooled at Prior to DNA isolation, the surface of each louse was 15 °C. PCR amplification was performed in a Peltier decontaminated to avoid external contamination, as de- PTC-200 model thermal cycler (MJ Research Inc., scribed previously [33]. Each louse specimen was cut in Watertown, MA, USA). Successful amplifications were half length-ways. DNA was extracted from one half and confirmed via electrophoresis on 1.5% agarose gel. Purifi- the remaining halves of the lice were frozen at -20 °C for cation of PCR products was performed using NucleoFast subsequent studies. DNA was extracted using the 96 PCR plates (Macherey Nagel EURL, Hoerdt, France) QIAamp DNA tissue extraction kit (Qiagen, Hilden, following the manufacturer’s instructions. The amplicons Germany) on the BioRobot EZ1 (Qiagen, Courtaboeuf, were sequenced using the Big Dye Terminator Cycle France) following the manufacturer’s instructions. DNA Sequencing Kit (Perkin Elmer Applied Biosystems, Foster was eluted in 100 μl of TE buffer and stored at -20 °C City, CA) with an ABI automated sequencer (Applied until the next investigation. Biosystems). The electropherograms obtained were then assembled and edited using ChromasPro software Genotypic status of lice (ChromasPro 1.7, Technelysium Pty Ltd., Tewantin, To identify the mitochondrial clades and to perform a Australia) and compared with those available in the phylogenetic study of the lice collected, all the DNA GenBank database by NCBI BLAST (http://blast.ncbi. samples were subjected to standard PCR, targeting a 347 nlm.nih.gov/Blast.cgi). bp fragment of the cytb gene. The reaction of amplifica- tion was conducted in a final volume of 50 μl, including Molecular screening for the presence of pathogens 25 μl Amplitaq gold master mix, 1 μl of each primer, 5 The DNA of all head lice were subject to amplification μl of DNA template, and water. The thermal cycling by qPCR using primers and probes targeting different condition was as follows: an initial denaturation step at genes of Rickettsia spp., Borrelia spp., B. quintana, Y. 147 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 4 of 11 Fig. 2 Map of head lice collection on Niger’s refugee (migrant population) and elementary schoolchildren (non-migrant population) from three localities in Algiers, Algeria pestis, Acinetobacter spp., C. burnetii and Anaplasma Acinetobacter spp. were subjected to standard PCR tar- spp., as previously reported (Table 1). All qPCRs were geting a 350 bp fragment of the rpoB gene (zone1) performed using a CFX96 Real-Time system (Bio-Rad, (Table 1). Negative and positive controls were included Marnes-la-Coquette, France) and the Eurogentec Master in each assay. Successful amplification was confirmed via Mix Probe PCR kit (Eurogentec, Liège, Belgium). We in- gel electrophoresis and amplicons were prepared and se- cluded target bacterial DNA as the positive control and quenced using similar methods as described for the cytb master mixtures as a negative control for each test. Sam- gene for lice above. ples were considered positive when the cycle threshold (Ct) was lower than 35 Ct. All C. burnetii positives sam- ples were confirmed by a second specific qPCR targeting Data analysis the IS30A spacer (Table 1). The head lice nucleotide sequences obtained in this In order to perform genotyping of C. burnetii, all posi- study were combined with the cytb database, compris- tive samples were subjected to PCR amplification of the ing haplotypes that span different geographical location multi-spacer typing (MST), targeting four intergenic in the five continents, as previously reported [9]. spacers (Cox2, Cox5, Cox18 and Cox22), as described MEGA 6.06 was used for the phylogenetic analyses previously [34]. To identify the species of the Acineto- under the Kimura 2-parameter model with 500 repli- bacter bacteria, all samples tested positive by qPCRs for cates, as described previously [35]. All obtained 148 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 5 of 11 Table 1 Oligonucleotide sequences of primers and probes used for real-time PCRs and conventional PCRs in this study Target Name Primers (5′-3′) and probes Source Pediculus humanus cytochrome b Cytb F_GAGCGACTGTAATTACTAATC [53] R_CAACAAAATTATCCGGGTCC Acinetobacter spp. RNA polymerase β subunit gene rpoB F_TACTCATATACCGAAAAGAAACGG [51] R_GGYTTACCAAGRCTATACTCAAC FAM-CGCGAAGATATCGGTCTSCAAGC-TAMRA rpoB F_TAYCGYAAAGAYTTGAAAGAAG [54] (zone1) R_CMACACCYTTGTTMCCRTGA Rickettsia spp. citrate synthase (gltA) RKNDO3 F_AATGCTCTTGCAGCTGGTTCT [55] R_TCGAGTGCTAATATTTTTGAAGCA FAM-CGGTGGTGTTAATGCTGCGTTACAACA-TAMRA Yersinia pestis plasminogen activator gene PLA F_ATGGAGCTTATACCGGAAAC [56] R_GCGATACTGGCCTGCAAG FAM-TCCCGAAAGGAGTGCGGGTAATAGG-TAMRA Borrelia spp. 16S ribosomal RNA Bor16S F_AGCCTTTAAAGCTTCGCTTGTAG [57] R_GCCTCCCGTAGGAGTCTGG FAM-CCGGCCTGAGAGGGTGAACGG-TAMRA Bartonella quintana Hypothetical intracellular effector 3-oxoacyl-synthase yopP F_TAAACCTCGGGGGAAGCAGA [58] gene R_TTTCGTCCTCAACCCCATCA FAM-CGTTGCCGACAAGACGTCCTTG-TAMRA fabF3 F_GCGGCCTTGCTCTTGATGA R_GCTACTCTGCGTGCCTTGGA FAM-TGCAGCAGGTGGAGAGAACGTG-TAMRA Anaplasma spp. 23S ribosomal RNA TtAna F_TGACAGCGTACCTTTTGCAT [59] R_TGGAGGACCGAACCTGTTAC FAM-GGATTAGACCCGAAACCAAG-TAMRA Coxiella burnetii Spacers IS1111 F_CAAGAAACGTATCGCTGTGGC [42] R_CACAGAGCCACCGTATGAATC FAM-CCGAGTTCGAAACAATGAGGGCTG-TAMRA IS30A F_ CGCTGACCTACAGAAATATGTCC R_ GGGGTAAGTAAATAATACCTTCTGG FAM-CATGAAGCGATTTATCAATACGTGTATGC- TAMRA Cox2 F_CAACCCTGAATACCCAAGGA [34] R_GAAGCTTCTGATAGGCGGGA Cox5 F_CAGGAGCAAGCTTGAATGCG R_TGGTATGACAACCCGTCATG Cox18 F_CGCAGACGAATTAGCCAATC R_TTCGATGATCCGATGGCCTT Cox22 F_GGGAATAAGAGAGTTAGCTCA R_CGCAAATTTCGGCACAGACC sequences of Acinetobacter species and C. burnetii were as described above, were used to determine the position analyzed using BLAST (www.ncbi.nlm.nih.gov/blast/ of Acinetobacter species identified in head lice of the Blast.cgi) and compared to sequences in the GenBank two population compared to another Acinetobacter database. Phylogenetic analyses using similar methods, available in the GenBank database. 149 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 6 of 11 Results genotyping of C. burnetii. We only succeeded in obtain- Lice clade and phylogenetic analysis ing sequences for the Cox22 spacer, probably due to Regarding the migrant population, we examined, in total, the low concentration of C. burnetii DNA in these head fifty female (≥ 24 years-old) and twenty child refugees (≤ 5 lice samples. Obtained sequences did not allow to iden- years-old). Twenty-eight of the fifty women (56%) and tify the C. burnetii genotype, but it corresponds to one three of the 20 (15%) children were infested with head lice of the eight possible MST genotypes of C. burnetii:8,9, and were included in this study. Thirty-seven head lice 10, 38, 43, 48, 50 and 53. Negative controls remained were collected from 31 individuals refugees from the negative in all PCR experiments. Nigerien refugee camp located in eastern Algiers, Algeria. The DNA of Acinetobacter spp. was detected in 20 Fourty-five head lice specimens were collected from out of 37 (54.05%) head lice collected from 25 out of 31 the non-migrant population. The sampling was per- (80.64%) Nigerien refugees and in 40 out of 45 (88.88%) formed in 5 elementary schools at 3 different locations head lice collected from 15 out of 27 (55.55%) school- in east Algiers, north Algeria. Of the 101 schholchildren children targeting the rpob gene using qPCR. As for the examined, 27 (26.73%) were infested by head lice and molecular identification of the Acinetobacter species, the totality were female aged between 6–11 years-old. we succeeded in amplifying a 350 bp fragment of the The highest infestation rate was recorded in the Bordj El rpoB (zone1) gene in all head lice positive in qPCR col- kiffan region with a rate of 48.14% (13/27), followed by lected from the two populations (migrant and the Bab Ezzouar and El Mohammadia regions, with a non-migrant). Based on a BLAST search, the compari- rate of 33.33% (9/27) and 18.51% (5/27), respectively. son of the nucleotide sequences with the GenBank In total, 82 head lice samples were collected from database sequences revealed the existence of one spe- Nigerien refugees and schoolchildren in eastern Algiers, cies of Acinetobacter for the head lice of Niger’srefu- and were then analyzed using PCR standard targeting gees sharing 99–100% identity with their corresponding the cytb gene to determine their clade. Results showed references and identified as A. baumannii. All lice that all head lice of the migrant population (37/82, belonged to the four haplotypes of clade E found in this 45.12%) were clade E, and defined the presence of four study (Fig. 3)(Table2). DNA of Acinetobacter spp. of different new haplotypes referred here as E52, E53, E54 head lice collected from schoolchildren were identified: and E55 (26/37, 70.27%) lice belonged to haplotype E52 24/40 (60%) positive-head lice as A. johnsonii, 12/40 (5/37, 13.51%) to haplotype E53 (4/37, 10.81%) to haplo- (30%) sequences as A. variabilis and 4/40 (10%) se- type E54 and (2/37, 5.40%) to haplotype E55. While the quences as A. baumannii,sharing99–100% identity head lice of the non-migrant population were clade A with their corresponding references and all belonged to and B, the analysis of the 45 cytb sequences yielded 34/ the two haplogroup A and B (Fig. 4). 82 lice (42.68%) belonged to the worldwide haplotype A5 within the clade A and 11/82 (13.41%) belonged to Discussion the B36 haplotype, the most widespread and prevalent In recent years, several countries around the world and within the B haplogroup (Table 2). These haplotypes, to- particularly those in the Horn of Africa, have endured gether with references from all the head lice and hap- humanitarian crisis conditions, including civil wars. The logroups, were used to construct a maximum likelihood European Union has received thousands of refugees and (ML) phylogenetic tree (Fig. 3). migrants, representing a risk factor for possible out- breaks of arthropod-borne diseases. The current refugee crisis in Europe has been characterized by the rapid Molecular detection of bacterial pathogens emergence of several cases of louse-borne relapsing fever In this study, the qPCR investigation of all lice samples caused by B. recurrentis which were diagnosed in the for Rickettsia spp., Borrelia spp., Y. pestis, B. quintana Netherlands, Germany, southern and northern Italy, and Anaplasma spp. produced no positive results. Belgium, Switzerland and Finland [28–32]. However, positive results were obtained when testing for Algeria is one of the countries in north Africa that has the presence of C. burnetii and Acinetobacter spp. also received thousands of refugees and migrants from The DNA of C. burnetii was detected in 3 out of 37 different countries, including Syria, Libya, Mali and (8.10%) of head lice of Nigerien refugees collected from Niger. An epidemiological survey was therefore estab- 3 of the 28 (10.71%) infested women but not reported lished in Algiers at a camp of Nigerien refugees that in head lice of schoolchildren. The positive lice for C. housed more than a hundred individuals crowded to- burnetii belonged to haplotypes E52 and E54 gether surviving in precarious conditions. These refugees (Fig. 3)(Table2). These results were confirmed by tar- have traveled thousands kilometers to reach their target geting two C. burnetii-specific genes using qPCR, sup- country, providing thus an ideal environment for the plemented by PCR-sequencing of one spacer for the spread of lice and the pathogens that they might carry. 150 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 7 of 11 Table 2 Detection of head lice clades and pathogens in the Nigerien refugees (migrant population) and schoolchildren (non-migrant population) in eastern Algiers, Algeria Location Population No. lice tested (%) Clade of lice (no.) % Haplotype (no.) % Pathogen (no.) % Bab Ezzouar Nigerien refugees 37 (45.12) E (37/37) 100 E52: (26/37) 70.27 Coxiella burnetii (3/37) 8.10 E53: (5/37) 13.51 Acinetobacter baumannii (20/37) 54.05 E54: (4/37) 10.81 E55: (2/37) 5.40 Schoolchildren 7 (8.53) A (3/7) 42.85 A5 (3/7) 42.85 Acinetobacter johnsonii (11/45) 24.44 B (4/7) 57.14 B36 (4/7) 57.14 Acinetobacter variabilis (3/45) 6.66 El Mohammadia Schoolchildren 11 (13.41) A (9/11) 81.81 A5 (9/11) 81.81 Acinetobacter johnsonii (7/45) 15.55 B (2/11) 18.18 B36 (2/11) 18.18 Acinetobacter variabilis (7/45) 15.55 Acinetobacter baumannii (4/45) 8.88 Bordj El kiffan Schoolchildren 27 (32.92) A (20/27) 74.07 A5 (20/27) 74.07 Acinetobacter johnsonii (6/45) 13.33 B (7/27) 25.92 B36 (7/27) 25.92 Acinetobacter variabilis (2/45) 4.44 Total 2 82 (100) A (34/82) 41.46 E52: (26/37) 70.27 Coxiella burnetii (3/82) 3.65 B (11/82) 13.41 E53: (5/37) 13.51 Acinetobacter baumannii (29/82) 35.36 E (37/82) 45.12 E54: (4/37) 10.81 Acinetobacter johnsonii (24/82) 29.26 E55: (2/37) 5.40 Acinetobacter variabilis (12/82) 14.63 A5 (34/82) 41.46 B36 (11/82) 13.41 Fig. 3 Maximum likelihood (ML) phylogram of the mitochondrial cytochrome b (cytb) gene. Phylogenetic inferences were conducted in MEGA 7 using the maximum likelihood method based on the Kimura 2-parameter. The mitochondrial clade memberships are indicated to the right of each tree. a Lice samples which are positive for Coxiella burnetii and Acinetobacter spp. and their genotypes (head or body lice) are specified (see legends at the top left). b Bacterial DNA detected in head lice reported in this study and the literature. The pathogenic bacteria in red are those naturally transmitted by body lice to humans 151 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 8 of 11 Fig. 4 Phylogenetic tree highlighting the position of the Acinetobacter species identified in head lice of Nigerien refuges and schoolchildren compared to another Acinetobacter available in the GenBank database. Phylogenetic inferences were conducted in MEGA 7 using the maximum likelihood method based on the Kimura 2-parameter model for nucleotide sequences. Statistical support for internal branches of the tree was evaluated by bootstrapping with 500 iterations Our study is the first to assess the genetic diversity as clade A and B, respectively. These data confirm the well as the occurrence of bacterial pathogens from head existence of clade A and B in Algeria, as reported by lice collected in camp of Nigerien refugees arriving from previous studies [36, 37, 27]. Zinder in Algeria, northern Africa. In this work, the This is the first report of clade E among head lice of pediculosis patients were female, 90.32% (28/31) were refugees coming from Niger, west Africa, to Algeria, as- adult and 9.67% (3/31) were girls. As mentioned earlier, suming that the head-louse infestation occurred in their we performed the same analysis for head lice specimens country of origin, and that clade E may be a vector for collected from schoolchildren of local population in the human pathogens. These results are not surprising, as order to have comparative sample to frame the results expected, they confirm that clade E has a west African obtained in this study. distribution, as reported by previous studies [9, 16]. The mitochondrial DNA analysis of the 37 head lice To date, human lice infestation remains a global public of the migrant population, and the 45 head lice of the health and social problem [38]. Although body lice are as- non-migrant population, showed the presence of three sumed to be the main vectors of pathogens, the epidemio- major haplogroups: A, B and E. All Niger’s refugees logical status of head lice as a vector of louse-borne head lice tested (37/82) belonged to clade E, including diseases is still misunderstood. Even though, it has been four different new haplotypes characterized in this demonstrated that the immune reactions of head lice to study: haplotype E52 was the most prevalent (70.27%), different pathogens are stronger than those of the body followed by haplotype E53 (13.51%), haplotype E54 louse, which may enable it to carry a large spectrum of (10.81%) and, finally, haplotype E55 (5.40%) (Table 2). pathogens [39, 40]. In natural settings, head lice belonging Previous studies have already reported that clade E is to different mitochondrial clades sampled from several limited to west African countries, namely Senegal and countries have been found to carry the DNA of several Mali [9]. It was then recently reported, for the first pathogen bacteria including B. quintana, B. recurrentis, B. time, in head lice from Bobigny (France), also sampled theileri, Acinetobacter spp. and Y. pestis,aswellasthe from families originating from west African countries DNA of C. burnetii, R. aeschlimannii and two potential [16]. The remaining head lice samples collected from new species of the genera Anaplasma and Ehrlichia,of sccholchildren in eastern Algiers, belonged to the unknown pathogenicity [11, 14, 16, 19, 21–27]. Experi- worldwide haplotype A5 (34/82, 42.68%) and the most mental studies have demonstrated that head lice may also widespread haplotype B6 (11/82, 13.41%), within the act as a vector for louse-borne diseases [33, 41]. 152 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 9 of 11 Coxiella burnetii, the causative agent of Q fever, is a was reported only in El Mohamammadia. Findings from highly infectious zoonotic intracellular bacterium. It is previous study on head lice collected from elementary found worldwide and has a diverse wide range of hosts schoolchildren in four localities in Algiers, Algeria have including mammals, birds, reptiles and arthropods, led to the same results, whither a widespreading infec- mainly ticks [42]. The infection is generally transmitted tion of head lice with the same Acinetobacter species to humans through direct contact (milk, urine, feces or found in our study [27]. semen from infected animals) as well as through aerosol Acinetobacter baumannii was isolated for the first time inhalation. It can be acute or chronic, exhibiting a wide from the body lice found on homeless people in range of clinical manifestations [42]. Q fever has been Marseille (France), as well as from various countries reported throughout the African continent with a higher around the world [50]. In recent years, the DNA of A. prevalence in western Africa, including several countries baumannii, in addition to several species of Acinetobac- such as Nigeria, Senegal, Ghana, Republic of Côte ter, were found in head lice collected from elementary d’Ivoire, Burkina Faso and Niger, representing a signifi- schoolchildren in Paris belonging to clade A [51], in cant public health threat [43]. In Niger, only one sero- Thailand belonging to A and C [14], in Algeria belong- logical study has so far been carried out on humans, in ing to clade A [27] and in head lice collected from Niamey, where 10% (17/177) of children aged between pygmy populations in the Republic of the Congo belong- one month and five years-old were found to be seroposi- ing to clades A, D, and C [11]. Acinetobacter baumannii tive for the C. burnetii infection [44]. Another study was DNA has also been detected in head lice collected from conducted on the prevalence of C. burnetii in animals of healthy individuals in Ethiopia [26]. The presence of A. the Maradi Region of Niger and showed that 32% (24/ baumannii DNA was even identified in ancient human 75) of goats that had experienced abortions were sero- head lice remains belonging to clade A [9]. Our study is positive, compared to 29% (12/75) of non-randomly se- the first to report the identification of A. baumannii lected goats without a history of abortion [45]. DNA in Nigerien head lice belonging to clade E, as well We found 8.10% of 37 head lice infected by C. burnetii as the identification of several species of Acinetobacter collected from three of 28 persons. This bacterium was in head lice belonging to clade B in Algeria. not reported in head lice sampled from the non-migrant Acinetobacter species are widespread in nature, includ- population. Coxiella burnetii was recently reported in ing in water, soil, living organisms and on the skin of pa- 1% of 600 head lice belonging to clade E infesting 5% of tients and healthy subjects [52]. Recent works have 117 individuals from Mali [21]. In contrast, the presence shown that the Acinetobacter infection can be highly of this bacterium was also investigated in Ethiopia on prevalent among body and head lice. However, it is still head and body lice, and results showed no evidence of not clear whether these Acinetobacter strains, present in C. burnetii in any of the specimens tested [46]. A field head lice, are the same as those responsible for human study in East Africa, showed that lice collected from in- infections [26], the clinical significance of this finding is dividuals, living in a place where an epidemic of Q fever still unknown. Furthermore, molecular evidence for the had occurred three months previously, could be spon- presence of DNA of C. burnetii and several Acinetobac- taneously infected with C. burnetii [47]. Another study ter species cannot distinguish between pathogens acci- also showed that, under experimental conditions, it is dentally acquired from the blood of infected individuals, possible to infect body lice with C. burnetii, although and those established in a competent vector which can human lice are not a known vector of this bacterium maintain and transmit the pathogen [17]. [48]. The presence of C. burnetii has never been re- ported in head lice belonging to the other clades; how- Conclusions ever, it has been recently reported for the firt time in 10/ Our study is the first to report that head lice from 524 (1.90%) body lice belonging to clade A collected Nigerian refugees belong to haplogroup E, and to from 2/19 (10.52%) homeless people in northern Algeria confirm that the clade E has a west African distribution. [49]. Based on our results and data from the literature, We also detected, for the first time, the presence of C. the role of human lice in the epidemiology of the C. bur- burnetii and A. baumannii in head lice from Niger. netii infection should be further investigated. Nevertheless, further studies of louse-borne pathogens This study also reveals the presence of A. baumannii are necessary to better understand the specificity of lice DNA in Nigerien refugees head lice, and the presence of to different pathogenic bacteria. Further studies are also A. johnsonii and A. variabilis in addition to A. bauman- required in order to explain their ability to harbor and, nii in head lice sampled from sccholchildren in eastern in certain cases, transmit pathogens from one person to Algiers, Algeria. The DNA of A. johnsonii and A. varia- another. Moreover, these results indicate that refugee bilis was reported in head lice collected from all three populations generate a potential risk of spreading emer- localities of the collection, while DNA of A. baumannii ging diseases and thus pose a significant epidemic threat 153 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 10 of 11 to public health in Algeria, where individual refugees liv- 5. Boutellis A, Abi-Rached L, Raoult D. The origin and distribution of human ing in conditions of poverty and promiscuity may serve lice in the world. Infect Genet Evol. 2014;23:209–17. 6. Brouqui P. Arthropod-borne diseases associated with political and social as indigenous reservoirs of multiple micro-organisms. disorder. Annu Rev Entomol. 2011;56:357–74. 7. Chosidow O. Scabies and pediculosis. Lancet Lond Engl. 2000;355:819–26. Abbreviations 8. Izri A, Uzzan B, Maigret M, Gordon MS, Bouges-Michel C. Clinical efficacy Cyt b: Cytochrome b; MST: Multispacer spacer typing; PCR: Polymerase chain and safety in head lice infection by Pediculus humanus capitis De Geer reaction; qPCR: Quantitative real-time polymerase chain reaction (Anoplura: Pediculidae) of a capillary spray containing a silicon-oil complex. Parasite. 2010;17:329–35. Acknowledgements 9. Amanzougaghene N, Mumcuoglu KY, Fenollar F, Alfi S, Yesilyurt G, Raoult D, We gratefully thank IHU Fondation Méditerranée-Infection for supporting the et al. High ancient genetic diversity of human lice, Pediculus humanus, from study. Israel reveals new insights into the origin of clade b lice. PLoS One. 2016;11: e0164659. Funding 10. Ashfaq M, Prosser S, Nasir S, Masood M, Ratnasingham S, Hebert PDN. High This study was supported by Méditerranée Infection and the National Research diversity and rapid diversification in the head louse, Pediculus humanus “ ’ ” Agency under the Investissements d avenir program, reference ANR-10-IAHU-03. (Pediculidae: Phthiraptera). Sci Rep. 2015;5:14188. 11. Amanzougaghene N, Akiana J, Mongo Ndombe G, Davoust B, Nsana NS, Availability of data and materials Parra HJ, et al. Head lice of pygmies reveal the presence of relapsing fever The data supporting the conclusions of this article are included within the borreliae in the Republic of Congo. PLoS Negl Trop Dis. 2016;10:e0005142. article. Representative sequences were submitted to the GenBank database 12. Drali R, Shako JC, Davoust B, Diatta G, Raoult D. A new clade of African under the accession numbers: MH392478, MH392479, MH392480 and MH392481. body and head lice infected by Bartonella quintana and Yersinia pestis - Democratic Republic of the Congo. Am J Trop Med Hyg. 2015;93:990–3. ’ Authors contributions 13. Light JE, Allen JM, Long LM, Carter TE, Barrow L, Raoult D, et al. Geographic Conceived and designed the experiments: OM, FF and IB. Collected samples: distributions and origins of human head lice (Pediculus humanus capitis) ML, MN and IB. Conducted the experiments: ML and OM. Analyzed the data: based on mitochondrial data. J Parasitol. 2008;94:1275–81. ML, NA and OM. Wrote the paper: ML, NA, MN, OM, FF and RD. All authors 14. Sunantaraporn S, Sanprasert V, Pengsakul T, Phumee A, Boonserm R, read and approved the final manuscript. Tawatsin A, et al. Molecular survey of the head louse Pediculus humanus capitis in Thailand and its potential role for transmitting Acinetobacter spp. Ethics approval and consent to participate Parasit Vectors. 2015;8:127. This study was approved by the Algerian Red Crescent organization (292/CRA/PRE), 15. Xiong H, Campelo D, Pollack RJ, Raoult D, Shao R, Alem M, et al. Second- Algeria. Head lice were collected from the scalps of Nigerien refugees generation sequencing of entire mitochondrial coding-regions (~15.4 kb) arriving in Algeria during a registered epidemiological study. Verbal consent holds promise for study of the phylogeny and taxonomy of human body was obtained from each infested individual. Written consent could not be lice and head lice. Med Vet Entomol. 2014;28:40–50. obtained because most of the subjects involved in the study were illiterate. The 16. Candy K, Amanzougaghene N, Izri A, Burn S, Durand R, Louni M, et al. anonymity of the individuals who provided the lice used in the present study Molecular survey of head and body lice, Pediculus humanus, in France. was preserved. Meanwhile, head lice samples from the local population were Vector Borne Zoonotic Dis. 2018;18:243–51. collected from the scalps of schholchildren under the approvement of the 17. Raoult D, Roux V. The body louse as a vector of reemerging human Ministry of National Education in Algeria (No. N14/0.0.2/224) with the parents’ diseases. Clin Infect Dis. 1999;29:888–911. explicit verbal consent. 18. Houhamdi L, Lepidi H, Drancourt M, Raoult D. Experimental model to evaluate the human body louse as a vector of plague. J Infect Dis. 2006;194: Competing interests 1589–96. The authors declare that they have no competing interests. 19. Piarroux R, Abedi AA, Shako JC, Kebela B, Karhemere S, Diatta G, et al. Plague epidemics and lice, Democratic Republic of the Congo. Emerg Infect Publisher’sNote Dis. 2013;19:505–6. Springer Nature remains neutral with regard to jurisdictional claims in published 20. Raoult D. A personal view of how paleomicrobiology aids our maps and institutional affiliations. understanding of the role of lice in plague pandemics. Microbiol Spectr. 2016;4. doi: https://doi.org/10.1128/microbiolspec.PoH-0001-2014. Author details 21. Amanzougaghene N, Florence F, Sangaré AK, Mahamadou S, Sissoko, 1IRD, AP-HM, SSA, VITROME, IHU-Méditerranée Infection, Aix Marseille Univ, Ogobara K, et al. Detection of bacterial pathogens including potential new Marseille, France. 2Laboratoire de Valorisation et Conservation des Ressources species in human head lice from human head lice from Mali. PLoS One. Biologiques (VALCOR), Faculté des Sciences, Université M’Hamed Bougara 2017;12:e0184621. Boumerdes, Boumerdes, Algeria. 3IRD, AP-HM, MEPHI, IHU-Méditerranée 22. Angelakis E, Rolain JM, Raoult D, Brouqui P. Bartonella quintana in head Infection, Aix-Marseille Univ, Marseille, France. 4Laboratoire Biodiversité et louse nits. FEMS Immunol Med Microbiol. 2011;62:244–6. Environnement: Interactions, Génomes, Département de Biologie, Université 23. Boutellis A, Veracx A, Angelakis E, Diatta G, Mediannikov O, Trape J-F, et al. des Sciences et Technologies Houari Boumediene, Bab Ezzouar, Algeria. Bartonella quintana in head lice from Senegal. Vector Borne Zoonotic Dis. 5Ecole Supérieure des Sciences de l’Aliment et des Industries 2012;12:564–7. Agro-Alimentaires, Algiers, Algeria. 24. Boutellis A, Mediannikov O, Bilcha KD, Ali J, Campelo D, Barker SC, et al. Borrelia recurrentis in head lice, Ethiopia. Emerg Infect Dis. 2013;19:796–8. Received: 21 February 2018 Accepted: 1 June 2018 25. Sangaré AK, Boutellis A, Drali R, Socolovschi C, Barker SC, Diatta G, et al. Detection of Bartonella quintana in African body and head lice. Am J Trop Med Hyg. 2014;91:294–301. References 26. Kempf M, Abdissa A, Diatta G, Trape JF, Angelakis E, Mediannikov O, et al. 1. Light JE, Smith VS, Allen JM, Durden LA, Reed DL. Evolutionary history of Detection of Acinetobacter baumannii in human head and body lice from mammalian sucking lice (Phthiraptera: Anoplura). BMC Evol Biol. 2010;10:292. Ethiopia and identification of new genotypes. Int J Infect Dis. 2012;16:14–6. 2. Reed DL, Smith VS, Hammond SL, Rogers AR, Clayton DH. Genetic analysis 27. Mana N, Louni M, Parola P, Bitam I. Human head lice and pubic lice reveal of lice supports direct contact between modern and archaic humans. PLoS the presence of several Acinetobacter species in Algiers, Algeria. Comp Biol. 2004;2:e340. Immunol Microbiol Infect Dis. 2017;53:33–9. 3. Veracx A, Raoult D. Biology and genetics of human head and body lice. 28. Antinori S, Mediannikov O, Corbellino M, Grande R, Parravicini C, Bestetti G, Trends Parasitol. 2012;28:563–71. et al. Louse-borne relapsing fever (Borrelia recurrentis) in a Somali refugee 4. Badiaga S, Brouqui P. Human louse-transmitted infectious diseases. Clin arriving in Italy: a re-emerging infection in Europe? PLoS Negl Trop Dis. Microbiol Infect. 2012;18:332–7. 2016;10:e0004522. 154 / 285 Louni et al. Parasites & Vectors (2018) 11:348 Page 11 of 11 29. Antinori S, Mediannikov O, Corbellino M, Raoult D. Louse-borne relapsing 54. La Scola B, Gundi VA, Khamis A, Raoult D. Sequencing of the rpoB gene and fever among East African refugees in Europe. Travel Med Infect Dis. 2016;14: flanking spacers for molecular identification of Acinetobacter species. Clin 110–4. Microbiol. 2006;44:827–32. 30. Cutler SJ. Refugee crisis and re-emergence of forgotten infections in 55. Rolain JM, Stuhl L, Maurin M, Raoult D. Evaluation of antibiotic susceptibilities Europe. Clin Microbiol Infect. 2016;22:8–9. of three rickettsial species including Rickettsia felis by a quantitative PCR DNA 31. Darcis G, Hayette MP, Bontems S, Sauvage AS, Meuris C, Van Esbroeck M, et assay. Antimicrob Agents Chemother. 2002;46:2747–51. al. Louse-borne relapsing fever in a refugee from Somalia arriving in 56. Nguyen-Hieu T, Aboudharam G, Signoli M, Rigeade C, Drancourt M, Raoult Belgium. J Travel Med. 2016;23 https://doi.org/10.1093/jtm/taw009. D. Evidence of a louse-borne outbreak involving typhus in Douai, 1710– 32. Hoch M, Wieser A, Löscher T, Margos G, Pürner F, Zühl J, et al. Louse-borne 1712 during the war of Spanish succession. PLoS One. 2010;5:e15405. relapsing fever (Borrelia recurrentis) diagnosed in 15 refugees from northeast 57. Parola P, Diatta G, Socolovschi C, Mediannikov O, Tall A, Bassene H, et al. Africa: Epidemiology and preventive control measures, Bavaria, Germany, Tick-borne relapsing fever borreliosis, rural senegal. Emerg Infect Dis. 2011; July to October 2015. Eurosurveillance. 2015;20 https://doi.org/10.2807/ 17:883–5. 1560-7917.ES.2015.20.42.30046. 58. Angelakis E, Socolovschi C. Raoult D. Short report: Bartonella quintana in 33. Robinson D, Leo N, Prociv P, Barker SC. Potential role of head lice, Pediculus Cimex hemipterus, Rwanda. Am J Trop Med Hyg. 2013;89:986–7. humanus capitis, as vectors of Rickettsia prowazekii. Parasitol Res. 2003;90: 59. Dahmani M, Davoust B, Benterki MS, Fenollar F, Raoult D, Mediannikov O. 209–11. Development of a new PCR-based assay to detect Anaplasmataceae and 34. Glazunova O, Roux V, Freylikman O, Sekeyova Z, Fournous G, Tyczka J, et al. the first report of Anaplasma phagocytophilum and Anaplasma platys in Coxiella burnetii genotyping. Emerg Infect Dis. 2005;11:1211–7. cattle from Algeria. Comp Immunol Microbiol Infect Dis. 2015;39:39–45. 35. Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: Molecular Evolutionary Genetics Analysis version 6.0. Mol Biol Evol. 2013;30:2725–9. 36. Boutellis A, Bitam I, Fekir K, Mana N, Raoult D. Evidence that clade A and clade B head lice live in sympatry and recombine in Algeria. Med Vet Entomol. 2015;29:94–8. 37. Drali R, Boutellis A, Raoult D, Rolain JM, Brouqui P. Distinguishing body lice from head lice by multiplex real-time PCR analysis of the Phum_ PHUM540560 gene. PLoS One. 2013;8:1–6. 38. Bonilla DL, Durden LA, Eremeeva ME, Dasch GA. The biology and taxonomy of head and body lice-implications for louse-borne disease prevention. PLoS Pathog. 2013;9:e1003724. 39. Kim JH, Previte DJ, Yoon KS, Murenzi E, Koehler JE, Pittendrigh BR, et al. Comparison of the proliferation and excretion of Bartonella quintana between body and head lice following oral challenge. Insect Mol Biol. 2017; 26:266–76. 40. Previte D, Olds BP, Yoon K, Sun W, Muir W, Paige KN, et al. Differential gene expression in laboratory strains of human head and body lice when challenged with Bartonella quintana, a pathogenic bacterium. Insect Mol Biol. 2014;23:244–54. 41. Murray ES, Torrey SB. Virulence of Rickettsia prowazekii for head lice. Ann N Y Acad Sci. 1975;266:25–34. 42. Mediannikov O, Fenollar F, Socolovschi C, Diatta G, Bassene H, Molez JF, et al. Coxiella burnetii in humans and ticks in rural Senegal. PLoS Negl Trop Dis. 2010;4:e654. 43. Vanderburg S, Rubach MP, Halliday JEB, Cleaveland S, Reddy EA, Crump JA. Epidemiology of Coxiella burnetii infection in Africa: a One Health systematic review. PLoS Negl Trop Dis. 2014;8:e2787. 44. Julvez J, Michault A, Kerdelhue C. Serological study of rickettsiose in Niamey, Niger. Med Trop (Mars). 1997;57:153–6 (In French). 45. Haumesser JB, Poutrel B. Rickettsiosis in the Niger. Epidemiological investigation carried out in the Maradi area. Rev Elev Med Vet Pays Trop. 1973;26:293–8. 46. Cutler S, Abdissa A, Adamu H, Tolosa T, Gashaw A. Bartonella quintana in Ethiopian lice. Comp Immunol Microbiol Infect Dis. 2012;35:17–21. 47. Giroud P, Jadin J. Infection latente et conservation de “Rickettsia burnetii” chez l’homme, le role du pou. Bull Soc Pathol Exot. 1954;47:764–5. 48. Babudieri B. Q fever: A zoonosis. Adv Vet Sci. 1959;5:82–182. 49. Louni M, Mana N, Bitam I, Dahmani M, Parola P, Fenollar F, et al. Body lice of homeless people reveal the presence of several emerging bacterial pathogens in northern Algeria. PLoS Negl Trop Dis. 2018;12:e0006397. 50. La Scola B, Raoult D. Acinetobacter baumannii in human body louse. Emerg Infect Dis. 2004;10:1671–3. 51. Bouvresse S, Socolovshi C, Berdjane Z, Durand R, Izri A, Raoult D, et al. No evidence of Bartonella quintana but detection of Acinetobacter baumannii in head lice from elementary schoolchildren in Paris. Comp Immunol Microbiol Infect Dis. 2011;34:475–7. 52. Vallenet D, Nordmann P, Barbe V, Poirel L, Mangenot S, Bataille E, et al. Comparative analysis of Acinetobacters: three genomes for three lifestyles. PLoS One. 2008;3:e1805. 53. Li W, Ortiz G, Fournier PE, Gimenez G, Reed DL, Pittendrigh B, et al. Genotyping of human lice suggests multiple emergences of body lice from local head louse populations. PLoS Negl Trop Dis. 2010;4:e641. 155 / 285 Article 8 : Molecular survey of head lice, Pediculus humanus capitis, in Democratic Republic of Congo En préparation pour soumission à PLOS Neglected Tropical Diseases 156 / 285 Molecular survey of head lice, Pediculus humanus capitis, in Democratic Republic of Congo Celia Scherelle BOUMBANDA KOYO1,4,5, Nadia AMANZOUGAGHENE2, Bernard DAVOUST2, Leon TSHILOLO3, Jean Bernard LEKANA-DOUKI4,6, Didier RAOULT2, Oleg MEDIANNIKOV2*, Florence FENOLLAR1 1Aix Marseille Univ, IRD, APHM, VITROME, IHU-Méditerranée Infection, Marseille, France 2Aix Marseille Univ, IRD, APHM, MEPHI, IHU-Méditerranée Infection, Marseille, France 3Monkole Mother and Child Hospital, Kinshasa, Democratic Republic of the Congo 4Unité d’Evolution, Epidémiologie et Résistances Parasitaires (UNEEREP), Centre International de Recherches Médicales de Franceville (CIRMF), B.P. 769 Franceville, Gabon 5Ecole Doctorale Régionale en Infectiologie Tropicale d’Afrique Centrale, B.P. 876 Franceville, Gabon 6Département de Parasitologie-Mycologie Médecine Tropicale, Faculté de Médecine, Université des Sciences de la Santé (USS), B.P. 4009 Libreville, Gabon *Corresponding author: Oleg Mediannikov Word abstract count: 239 Word text count: 2,847 Keywords: Head lice; clade E; Acinetobacter baumannii; Acinetobacter spp.; Democratic Republic of Congo 157 / 285 Abstract Head louse, Pediculus humanus capitis, is obligatory blood-sucking ectoparasite, distributed worldwide. Phylogenetically, it occurs in five divergent mitochondrial clades (A- E); each exhibiting a particular geographical distribution. Recent studies suggest that, as in the case of body louse, head louse could be a disease vector. We aimed to study the genetic diversity of head lice collected from Democratic Republic of Congo (DR Congo) and screen for louse-borne pathogens. Among 181 head lice, collected from 27 individuals, 67.4% (122/181) were clade A and 24.3% (44/181) clade D. Besides, for the first time in this area, we found clade E in 8.3% (15/181) of tested lice, from two infested individuals. Dual infestation with clades A and D was observed for 44.4% individuals. Regarding louse-borne bacteria, we looked for Bartonella quintana, B. recurrentis, Rickettsia prowazekii, Anaplasma spp., Yersinia pestis, Coxiella burnetii, and Acinetobacter spp. Only, widespread infection of head lice with several species of Acinetobacter was found. Overall, six species were detected in 18.2% of tested lice infesting 59.3% individuals. Acinetobacter baumannii (8.29%) was the most prevalent followed by Acinetobacter johnsonii (1.66%), Acinetobacter soli (1.66%), Acinetobacter pittii (1.66%), and Acinetobacter guillouaie (1.10%). One potential new species Candidatus Acinetobacter pediculi was also identified. For the first time, the presence of clade E head lice is observed in DR Congo. This also the first report of the presence of Acinetobacter species DNAs in human head lice in DR Congo. 158 / 285 INTRODUCTION Lice are obligatory hematophagous ectoparasites of placental mammals, including humans [1]. More than 550 species of lice have been identified worldwide; each of them is specific to a mammal host species [2]. Two lice species infested humans: Pediculus humanus and Pthirus pubis [2]. The former is of great public health concern and includes two ecotypes: head lice Pediculus humanus capitis, which live in the scalp region, and body lice Pediculus humanus humanus which live in clothing [2,3]. Studies based on mitochondrial genes appear to separate head and body lice into five divergent clades (A, B, C, D, and E) exhibiting some geographic differences [4–6]. Head lice encompass all diversity while body lice belong only to clades, A and D [4,7]. Clade A has a worldwide distribution [2,8]. Clade B was first found in North and Central America, Europe and Australia, and most recently found in North and Central Africa [4,9]. This clade was also reported at Israel, on head lice remains dating approximately to 2,000 year-old [5]. Clade C is found in Africa and Asian continents, and recently in France [7,10]. Clade D was only found in central Africa [6,8]. Lastly, clade E consists of head lice from West Africa (Senegal and Mali) [5] and was recently detected among head lice in France (Candy et al, 2018). Based on archeological remains, the Pediculus louse is thought to be an ancient parasite that had long association with their human hosts [11]. Because of this long association lice have become a model for study of cophylogenetic relationship between hosts and parasites [12]. Besides their role of pests, lice are disease vectors, more importantly body lice which is the main vector of at least three pathogenic bacteria that are killed millions of people, namely: Rickettsia prowazekii, the causative agent of epidemic typhus, Borrelia recurrentis, the causative agent of relapsing fever, and Bartonella quintana the causative agent of trench fever [13]. There are natural and experimental observations showing that body lice can also transmit 159 / 285 Yersinia pestis, the causative agent of plague, and that they may be the pandemic vector of this agent [14,15]. Although body lice are currently assumed to be more potent vectors of pathogens, the potential role of head lice as vector, is not fully understood. Studies demonstrated that immune reactions of head lice to different pathogens are stronger than those of body lice which obviously may carry a broad spectrum of pathogens [16,17]. Indeed, several field studies reports finding of the body louse-borne pathogens on head lice collected worldwide. This is the case of B. quintana, B. recurrentis, Borrelia theileri, Coxiella burnetii, and Y. pestis DNAs detected in head lice belonging to different mitochondrial clades [6–8,18–20]. Several Acinetobacter spp. were also detected in human head lice [8,21,22]. Furthermore, experimental infections with R. prowazekii have shown that head lice can be readily infected and disseminate these pathogens in their feces, demonstrating that these lice have the potential to be a vector pathogen under optimal epidemiologic conditions [23]. This fact poses a very substantial health risk to infested persons because such infestations are very prevalent worldwide and the epidemics still occurring regularly. Children are at increased risk, regardless of hygiene conditions and social status [24]. In this study, we aimed to investigate genetic diversity in head lice collected in Democratic Republic of Congo (DR Congo) and look for pathogenic bacteria in these lice. MATERIAL AND METHODS Ethics statement and louse sampling This study was approved by the central ethic committee (CEL) of CEFA (CEntre de Formation et d’Appui sanitaire) associated with Monkole Hospital Center (N/Réf:020/CEFA- MONKOLE/CEL/2017). Lice collection was performed at medical center of Monkole located at Kinshasa, largest city and capital of the DR Congo. Only patients who given their informed consent were included in this study. In total 27 patients were enrolled and thoroughly examined 160 / 285 for detection of both body and head lice. They were coming from 11 localities very closed geographically. A total of 181 head lice were collected from these patients. No body lice were found during the examination. Collected lice were preserved in 70% alcohol, before to be sent in our laboratory in Marseille (France), at room temperature. DNA extraction To avoid false-positive linked to bacteria of lice external surface, each louse specimen was decontaminated as described previously [25]. Then, each louse was dried and cut in half lengthwise. Half was frozen at -20°C for later use. The other was crushed in sterile Eppendorf tube; total-DNA was extracted using a DNA extraction kit, QIAamp Tissue Kit (Qiagen®, Courtaboeuf, France) using the EZ1 apparatus following the manufacturer’s protocol. The DNA was eluted in 100 μl of TE (10/1) buffer and stored at +4°C until use for PCR amplifications. DNA quantity and quality were assessed using a NanoDrop ND-1000 (Thermo Fisher Scientific®, Waltham, MA, USA). Genotypic status of lice Identification of louse mitochondrial clade by qPCR assays. To identify the mitochondrial clades of the lice included in this study, all DNA samples were analyzed using clade-specific quantitative real-time PCR (qPCR) assays that targeted a portion of cytochrome b (cytb) gene specific to each of the five clades, as previously described [7]. We used lice with known clades as positive controls and master mixtures as a negative control for each assay. All PCR amplifications were carried out using a CFX96 Real-Time system (Bio-Rad® Laboratories, Foster City, CA, USA) as previously described [7]. Sequences of primers and probes are shown in Table 1. Cytochrome b amplification and haplotype determination. For phylogenetic study, DNA samples from 54 head lice randomly selected from the total number of lice, were subjected to standard PCR targeting a 347-bp fragment of the Cytb gene using the primers and 161 / 285 conditions as previously described [26]. PCRs consisted of 50 µl volume including 25 µl Amplitaq gold master mix, 1 µl of each primer, 5 μl of DNA template, and water. The thermal cycling profile was one incubation step at 95°C for 15 minutes, 40 cycles of one minute at 95°C, 30 seconds at 56°C and one minute at 72°C, followed by a final extension for five minutes at 72°C. PCR amplification was performed in a Peltier PTC-200 model thermal cycler (MJ Research Inc®. Watertown, MA, USA). The success of amplification was confirmed by electrophoresis on agarose gel. Purification of PCR products was performed using NucleoFast 96 PCR plates (Macherey-Nagel EURL®, Hoerdt, France) according to the manufacturer’s instructions. The amplicons were sequenced using the Big Dye Terminator Cycle Sequencing Kit (Perkin Elmer Applied Biosystems®, Foster City, CA) with an ABI automated sequencer (Applied Biosystems). The electrophoregrams which were obtained were assembled and edited using ChromasPro software (ChromasPro 1.7, Technelysium Pty Ltd., Tewantin, Australia). Molecular screening for the presence of bacterial DNA The qPCRs were performed to screen all lice samples, using previously reported primers and probes, for Borrelia spp., B. quintana, Acinetobacter spp., Rickettsia spp, R. prowazekii, Y. pestis, Anaplasma spp., and C. burnetii. Sequences of primers and probes used in this study for qPCRs and standard PCRs are showed in Table 1. qPCRs were performed using a CFX96 Real-Time system (Bio-Rad®) and the Roche LightCycler® 480 Probes Master Mix PCR kit (Roche Applied Science, Mannheim Germany). We included DNA extracts of the targeted bacteria as positive controls and master mixtures as negative control for each assay. We considered samples to be positive when the cycle’s threshold (Ct) was lower than 35 Ct [27]. To identify the species of Acinetobacter, all positive samples from qPCR were subjected to standard PCR, targeting a portion of the rpoB gene (zone 1) using primers and conditions 162 / 285 previously described [28]. Successful amplification was confirmed via gel electrophoresis and amplicons were prepared and sequenced using similar methods as described for the cytb gene for lice above. Data analysis To obtain head lice cytb sequences, unique haplotypes were defined using DnaSPv5.10 and compared with the cytb haplotypes and combined cytb haplotypes reported by Amanzougaghene et al. [8]. In order to investigate the possible relationships between the haplotypes, the median-joining (MJ) network using the method of Bandelt was constructed with the program NETWORK4.6 (www.fluxus-engineering.com/sharenet.htm) [29]. Phylogenetic analyses and tree reconstruction were performed using MEGA software version 6.06 [30]. All obtained sequences of Acinetobacter species were analyzed using BLAST (www.ncbi.nlm.nih.gov/blast/Blast.cgi) and compared with sequences in GenBank database. A maximum-likelihood methods was used to infer the phylogenetic analyses and tree reconstruction was performed using MEGA software version 6.06 [30]. RESULTS Lice clade and phylogenetic analyses Overall, 181 head lice were collected from 27 obviously healthy individuals, all were female. The average number of lice by person was 6.70±6.61. No impairment of general condition was observed during the examination. Nevertheless, some person presented signs such as itching, tickling on the neck or the head; only one person had headaches. All collected lice were tested by qPCRs to determine their clade. Our result showed that 67.4% (122/181) lice belonged to clade A, 24.3% (44/181) lice to clade D, and only 8.3% (15/181) lice to clade E. Among the 27 persons, 15 (55.56%) were mono-infested by one clade of lice. Among them, 8 (29.63%) individuals were only infested with clade A lice, 5 (18.52%) 163 / 285 only with clade D lice, and 2 (7.41%) only with clade E lice. Finally, dual infestation was observed in 12 individuals (44.44%), and only with clades A and D (Table 2). The analysis of 54 cytb sequences yielded 43 variable positions defining 10 different haplotypes. One haplotype belonged to the worldwide haplotype A5 within clade A. Four haplotypes also belonged to clade A, were new and named here as A66-A69 (Table 3). Within clade D, four haplotypes were identified; one belonged to the D60 haplotype. The other three haplotypes were novel and are referred here as D74-D76. The remaining haplotype belonged to clade E, was novel and referred here as E62 (Table 3). Phylogenetic tree and median-joining analyses result in similar; all the cytb sequences were divided among five major supported clades, represented by five connected subnetworks distinct groups as show in the MJ network corresponding to the known clades: A, D, B, C, and E. The 10 haplotypes of our study fell into the three clades A, D, and E (Figures 1 and 2). Molecular detection of bacterial DNA All the head lice tested on qPCRs were found negative for B. quintana, Y. pestis, C. burnetii, Borrelia, Anaplasma, Rickettsia spp., and R. prowazekii. DNA of Acinetobacter spp was detected in 33/181 (18.23%) of the tested head lice, infesting 16/27 (59.26%) individuals. Sequencing of 340-bps fragment rpoB gene coupled with blast analysis identified six Acinetobacter species, including one potential new one(Table 4). The most prevalent species was A. baumannii with a prevalence of 8.29% (15/181). A. baumannii was detected in 7 lice from clade A, 6 from clade D, and 2 from clade E. Then, we detected A. pittii, A. soli, and A. johnsonii, with the same prevalence of 1.66% (3/181). Finally, A. guillouiae was found in 1.1% (2/181) of tested lice. Among the 3 lice positive for A. pittii, 2 belonged to clade A and 1 to clade D. The same distribution was observed for A. johnsonii. The 3 lice positive for A. soli belonged to clade A. One positive louse for A. guillouiae belonged to clade E, the other to clade A. All these species sharing 99-100% identity with their 164 / 285 corresponding reference Acinetobacter species. The potential new species showed prevalence of 1.1% (2/181); it is named here as Candidatus Acinetobacter pediculi. The most closely- related species is: A. guillouiae (GenBank number FJ754439) with 94.9% similarity (337 of 355 base positions in common). The 2 positive lice for the undescribed species of Acinetobacter (Candidatus Acinetobacter pediculi) belonged to clade A. The 5 other ones presented also some similarities with Acinetobacter. However, the sequences were of poor quality. The distribution of Acinetobacter species according to lice clades are presented in Table 4. The phylogenetic position of A. baumannii, A. pittii, A. soli, A. johnsonii and A. guillouiae is also given in Figure 3. The partial rpoB sequences of the Acinetobacter species obtained in this study were deposited in the GenBank under the accession number: From MH230910 to MH230920. DISCUSSION Hundreds of millions of head louse pediculosis occur every year worldwide, mostly among children of 3 to 14 year-old [31]. The increased resistance in pediculicides is, on one hand, responsible of the globally increase of head lice infestation [32], and the most threat is that pathogenic bacteria transmitted at baseline by body lice, had been found in head lice [7]. This brings us to wonder about the vectorial role of the latter. Here, we studied the genetic diversity of head lice collected from DR Congo. The presence of lice from clades A, D, and E was observed. The most prevalent clade was A , confirming its worldwide distribution, followed by clade D , and by clade E. Clades A and D were already reported in this area, but this is the first report of clade E in Central Africa, which is more abundant in West Africa [6]7,[33]. All positive lice for clade E arise from only 2 people. Several hypotheses could be suggested such as the recent arrival of these people from West African country, a close contact with West African populations, or a previous implantation of these lice but with low prevalence. However, only one haplotype for all clade E lice was observed. Besides, it was a new one, called here E62, and never described in West Africa. 165 / 285 Overall, these data show that the current repartition of human lice clade is not definitive. Increasing the samples sizes and extending the geographic coverage are needed to better determine the intra and interclade diversity [4]. In addition to the inter-haplogroup diversity, P. humanus also shown intra-haplogroup diversity, which is illustrated by many distinct A, B, D, E, and C haplotypes [4,8,10,34], results supported by our findings. Indeed, among the 54 head lice cytb sequences analyzed, 10 different haplotypes were identified; in which 8 haplotypes were novel. There are several reports that states that co-infestation by different mtDNA clades of human lice in the same individual can occurs and it was found to be associated with clades A and B [9,35], clades A and C [35,36], and also for clades A and D [6], suggesting that these different clades can live in sympatry and interbreed [9,36]. In our study, only half of the people were mono-infested by one clade of lice. Dual infestation was only observed with clades A and D in 12 individuals (44%). This data was consistent with previous report from Drali et al., which found dual infestation with clades A and D among 14 of 37 (37.8%) infested people in DR Congo [6]. It would be interesting to know whether or not any evidence of gene exchange and recombination events occur between these different clades or if lice living in sympatry. Nevertheless, there was none dual infestation between clades A and E, and between clades D and E. It once was believed that, only body lice transmitted bacterial pathogens to infested persons, currently the vectorial role of head lice is discussed. We screened all the 181 head lice collected for the presence of the DNA of several bacterial pathogens. Several epidemiological studies have reported the presence of the bacterial DNA on head lice collected from different parts of the world. In our study only, Acinetobacter species was found with A. baumannii the most prevalent consistence with previous studies that shown that A. baumannii is the more abundant species found in head and body lice [22]. Another study performed in Congo 166 / 285 (Brazzaville) among the pygmies populations have found 10.39% of A. baumannii, as well as a large scale of other Acinetobacter species such as: Acinetobacter junii (18.31%), Acinetobacter ursingii (14.35%), Acinetobacter johnsonii (9.40%), Acinetobacter schindleri (8.41%), Acinetobacter nosocomialis (3.18%), Acinetobacter lwoffii (4.45%), and Acinetobacter towneri (1.98%) [8]. Among Acinetobacter species, A. baumannii, is the most important species, observed worldwide, and involved in hospital acquired infections, including epidemics that are a real challenge for public health. Currently, A. baumannii is recognized as a pathogen responsible for nosocomial infection, but also community-acquired infections and war- and natural disaster-related infections, such as war wounds in troops from Iraq and Afghanistan [37–39][40]. The more interesting is the discovery of one potential new species Candidatus Acinetobacter pediculi. Our study is the first to describe A. soli, A. pittii, and A. guillouiae in human lice. Unlike to A. guillouiae which is an environmental species, scarcely associated with human infection, A. soli and A. pitti has been isolated from clinical samples and are associated to carbapenem resistance [41,42]. The presence of B. quintana and Y. pestis has been already reported in DR Congo in head and body lice and a possible exchange of pathogens between head and body lice has been suggested [43]. In conclusion, we highlighted the presence of clade E head lice in Central Africa. The more prevalent head lice clades in DR Congo were clades A and D. Several Acinetobacter species were detected, including one potential new one. More attention is now paid to reservoirs of these ubiquitous opportunistic bacteria and their potential involvement in human infections. 167 / 285 Acknowledgments This study was supported by Méditerranée Infection and the National Research Agency under the program « Investissements d’avenir », reference ANR-10-IAHU-03. CSBK is PhD student fund by Gabonese Governement. We thank medical staff involved in this study and patients who agree to participated in this study. 168 / 285 References 1. Barker SC. Phylogeny and classification, origins, and evolution of host associations of lice. Int J Parasitol. 1994;24: 1285–1291. doi.org/10.1016/0020-7519(94)90195-3 2. Bonilla DL, Durden LA, Eremeeva ME, Dasch GA. The Biology and Taxonomy of Head and Body Lice—Implications for Louse-Borne Disease Prevention. PLoS Pathog. 2013;9. doi:10.1371/journal.ppat.1003724 3. Veracx A, Raoult D. Biology and genetics of human head and body lice. Trends Parasitol. 2012;28: 563–571. doi:10.1016/j.pt.2012.09.003 4. Ashfaq M, Prosser S, Nasir S, Masood M, Ratnasingham S, Hebert PDN. High diversity and rapid diversification in the head louse, Pediculus humanus (Pediculidae: Phthiraptera). Sci Rep. 2015;5. doi:10.1038/srep14188 5. Amanzougaghene N, Mumcuoglu KY, Fenollar F, Alfi S, Yesilyurt G, Raoult D, et al. High Ancient Genetic Diversity of Human Lice, Pediculus humanus, from Israel Reveals New Insights into the Origin of Clade B Lice. PloS One. 2016;11: e0164659. doi:10.1371/journal.pone.0164659 6. Drali R, Shako J-C, Davoust B, Diatta G, Raoult D. A New Clade of African Body and Head Lice Infected by Bartonella quintana and Yersinia pestis-Democratic Republic of the Congo. Am J Trop Med Hyg. 2015;93: 990–993. doi:10.4269/ajtmh.14-0686 7. Amanzougaghene N, Fenollar F, Sangaré AK, Sissoko MS, Doumbo OK, Raoult D, et al. Detection of bacterial pathogens including potential new species in human head lice from Mali. PLOS ONE. 2017;12: e0184621. doi:10.1371/journal.pone.0184621 8. Amanzougaghene N, Akiana J, Mongo Ndombe G, Davoust B, Nsana NS, Parra H-J, et al. Head Lice of Pygmies Reveal the Presence of Relapsing Fever Borreliae in the Republic of Congo. PLoS Negl Trop Dis. 2016;10: e0005142. doi:10.1371/journal.pntd.0005142 9. Boutellis A, Bitam I, Fekir K, Mana N, Raoult D. Evidence that clade A and clade B head lice live in sympatry and recombine in Algeria. Med Vet Entomol. 2015;29: 94–98. doi:10.1111/mve.12058 10. Light JE, Allen JM, Long LM, Carter TE, Barrow L, Suren G, et al. Geographic distributions and origins of human head lice (Pediculus humanus capitis) based on mitochondrial data. J Parasitol. 2008;94: 1275–1281. doi:10.1645/GE-1618.1 11. Ascunce MS, Toups MA, Kassu G, Fane J, Scholl K, Reed DL. Nuclear Genetic Diversity in Human Lice (Pediculus humanus) Reveals Continental Differences and High Inbreeding among Worldwide Populations. PLOS ONE. 2013;8: e57619. doi:10.1371/journal.pone.0057619 169 / 285 12. Demastes JW, Spradling TA, Hafner MS, Spies GR, Hafner DJ, Light JE. Cophylogeny on a fine scale: Geomydoecus chewing lice and their pocket gopher hosts, Pappogeomys bulleri. J Parasitol. 2012;98: 262–270. doi:10.1645/GE-2904.1 13. Raoult D, Roux V. The body louse as a vector of reemerging human diseases. Clin Infect Dis Off Publ Infect Dis Soc Am. 1999;29: 888–911. doi:10.1086/520454 14. Houhamdi L, Lepidi H, Drancourt M, Raoult D. Experimental model to evaluate the human body louse as a vector of plague. J Infect Dis. 2006;194: 1589–1596. doi:10.1086/508995 15. Raoult D. A Personal View of How Paleomicrobiology Aids Our Understanding of the Role of Lice in Plague Pandemics. Microbiol Spectr. 2016;4. doi:10.1128/microbiolspec.PoH- 0001-2014 16. Kim JH, Previte DJ, Yoon KS, Murenzi E, Koehler JE, Pittendrigh BR, et al. Comparison of the proliferation and excretion of Bartonella quintana between body and head lice following oral challenge. Insect Mol Biol. 2017;26: 266–276. doi:10.1111/imb.12292 17. Previte D, Olds BP, Yoon K, Sun W, Muir W, Paige KN, et al. Differential gene expression in laboratory strains of human head and body lice when challenged with Bartonella quintana, a pathogenic bacterium. Insect Mol Biol. 2014;23: 244–254. doi:10.1111/imb.12077 18. Angelakis E, Diatta G, Abdissa A, Trape J-F, Mediannikov O, Richet H, et al. Altitude- dependent Bartonella quintana genotype C in head lice, Ethiopia. Emerg Infect Dis. 2011;17: 2357–2359. doi:10.3201/eid1712.110453 19. Boutellis A, Veracx A, Angelakis E, Diatta G, Mediannikov O, Trape J-F, et al. Bartonella quintana in head lice from Sénégal. Vector Borne Zoonotic Dis Larchmt N. 2012;12: 564– 567. doi:10.1089/vbz.2011.0845 20. Boutellis A, Mediannikov O, Bilcha KD, Ali J, Campelo D, Barker SC, et al. Borrelia recurrentis in head lice, Ethiopia. Emerg Infect Dis. 2013;19: 796–798. doi:10.3201/eid1905.121480 21. Bouvresse S, Socolovshi C, Berdjane Z, Durand R, Izri A, Raoult D, et al. No evidence of Bartonella quintana but detection of Acinetobacter baumannii in head lice from elementary schoolchildren in Paris. Comp Immunol Microbiol Infect Dis. 2011;34: 475– 477. doi:10.1016/j.cimid.2011.08.007 22. Sunantaraporn S, Sanprasert V, Pengsakul T, Phumee A, Boonserm R, Tawatsin A, et al. Molecular survey of the head louse Pediculus humanus capitis in Thailand and its potential role for transmitting Acinetobacter spp. Parasit Vectors. 2015;8: 127. doi:10.1186/s13071-015-0742-4 170 / 285 23. Robinson D, Leo N, Prociv P, Barker SC. Potential role of head lice, Pediculus humanus capitis, as vectors of Rickettsia prowazekii. Parasitol Res. 2003;90: 209–211. doi:10.1007/s00436-003-0842-5 24. Chosidow O. Scabies and pediculosis. Lancet Lond Engl. 2000;355: 819–826. 25. La Scola B, Fournier P-E, Brouqui P, Raoult D. Detection and Culture of Bartonella quintana, Serratia marcescens, and Acinetobacter spp. from Decontaminated Human Body Lice. J Clin Microbiol. 2001;39: 1707–1709. doi:10.1128/JCM.39.5.1707-1709.2001 26. Li W, Ortiz G, Fournier P-E, Gimenez G, Reed DL, Pittendrigh B, et al. Genotyping of human lice suggests multiple emergencies of body lice from local head louse populations. PLoS Negl Trop Dis. 2010;4: e641. doi:10.1371/journal.pntd.0000641 27. Sokhna C, Mediannikov O, Fenollar F, Bassene H, Diatta G, Tall A, et al. Point-of-Care Laboratory of Pathogen Diagnosis in Rural Senegal. PLoS Negl Trop Dis. 2013;7: e1999. doi:10.1371/journal.pntd.0001999 28. La Scola B, Gundi VAKB, Khamis A, Raoult D. Sequencing of the rpoB gene and flanking spacers for molecular identification of Acinetobacter species. J Clin Microbiol. 2006;44: 827–832. doi:10.1128/JCM.44.3.827-832.2006 29. Bandelt HJ, Forster P, Röhl A. Median-joining networks for inferring intraspecific phylogenies. Mol Biol Evol. 1999;16: 37–48. doi:10.1093/oxfordjournals.molbev.a026036 30. Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: Molecular Evolutionary Genetics Analysis Version 6.0. Mol Biol Evol. 2013;30: 2725–2729. doi:10.1093/molbev/mst197 31. Gratz NG, Scheme WHOD of C of TDWPE. Human li e : thei p e ale e, o t ol a d esista e to i se ti ides : a e ie 1 -1997. 1997; Available: http://www.who.int/iris/handle/10665/63791 32. Burgess IF. Human Lice and Their Control. Annu Rev Entomol. 2004;49: 457–481. doi:10.1146/annurev.ento.49.061802.123253 33. Raoult D, Reed DL, Dittmar K, Kirchman JJ, Rolain J-M, Guillen S, et al. Molecular Identification of Lice from Pre-Columbian Mummies. J Infect Dis. 2008;197: 535–543. doi:10.1086/526520 34. Ascunce MS, Fane J, Kassu G, Toloza AC, Picollo MI, González-Oliver A, et al. Mitochondrial diversity in human head louse populations across the Americas. Am J Phys Anthropol. 2013;152: 118–129. doi:10.1002/ajpa.22336 35. Xiong H, Campelo D, Boutellis A, Raoult D, Alem M, Ali J, et al. SNPs in Entire Mito ho d ial Ge o e Se ue es ≈1 . k a d ox1 Se ue es ≈ p Resol e 171 / 285 Body and Head Lice From Doubly Infected People From Ethiopia, China, Nepal, and Iran But Not France. J Med Entomol. 2014;51: 1199–1207. doi:10.1603/ME14001 36. Veracx A, Boutellis A, Raoult D. Genetic recombination events between sympatric Clade A and Clade C lice in Africa. J Med Entomol. 2013;50: 1165–1168. 37. Eveillard M, Kempf M, Belmonte O, Pailhoriès H, Joly-Guillou M-L. Reservoirs of Acinetobacter baumannii outside the hospital and potential involvement in emerging human community-acquired infections. Int J Infect Dis. 2013;17: e802–e805. doi:10.1016/j.ijid.2013.03.021 38. Peleg AY, Seifert H, Paterson DL. Acinetobacter baumannii: Emergence of a Successful Pathogen. Clin Microbiol Rev. 2008;21: 538–582. doi:10.1128/CMR.00058-07 39. Johanson WG, Pierce AK, Sanford JP. Changing pharyngeal bacterial flora of hospitalized patients. Emergence of gram-negative bacilli. N Engl J Med. 1969;281: 1137–1140. doi:10.1056/NEJM196911202812101 40. Abraham EP, Chain E. An enzyme from bacteria able to destroy penicillin. 1940. Rev Infect Dis. 1988;10: 677–678. 41. Pellegrino FLPC, Vieira VV, Baio PVP, dos Santos RMR, dos Santos ALA, de Barros Santos NG, et al. Acinetobacter soli as a Cause of Bloodstream Infection in a Neonatal Intensive Care Unit▿ . J Clin Microbiol. 2011;49: 2283–2285. doi:10.1128/JCM.00326-11 42. Endo S, Sasano M, Yano H, Arai K, Aoyagi T, Hatta M, et al. First Carbapenem-Resistant Isolates of Acinetobacter soli in Japan. Antimicrob Agents Chemother. 2012;56: 2786– 2787. doi:10.1128/AAC.00021-12 43. Piarroux R, Abedi AA, Shako J-C, Kebela B, Karhemere S, Diatta G, et al. Plague epidemics and lice, Democratic Republic of the Congo. Emerg Infect Dis. 2013;19: 505–506. 44. Nguyen-Hieu T, Aboudharam G, Signoli M, Rigeade C, Drancourt M, Raoult D. Evidence of a louse-borne outbreak involving typhus in Douai, 1710-1712 during the war of Spanish succession. PloS One. 2010;5: e15405. doi:10.1371/journal.pone.0015405 45. Parola P, Diatta G, Socolovschi C, Mediannikov O, Tall A, Bassene H, et al. Tick-borne relapsing fever borreliosis, rural senegal. Emerg Infect Dis. 2011;17: 883–885. doi:10.3201/eid1705.100573 46. Dahmani M, Davoust B, Rousseau F, Raoult D, Fenollar F, Mediannikov O. Natural Anaplasmataceae infection in Rhipicephalus bursa ticks collected from sheep in the French Basque Country. Ticks Tick-Borne Dis. 2017;8: 18–24. doi:10.1016/j.ttbdis.2016.09.009 47. Mediannikov O, Diatta G, Fenollar F, Sokhna C, Trape J-F, Raoult D. Tick-borne rickettsioses, neglected emerging diseases in rural Senegal. PLoS Negl Trop Dis. 2010;4. doi:10.1371/journal.pntd.0000821 172 / 285 Figure 1. Maximum-likelihood (ML) phylogram of the mitochondrial Cytb haplotypes. A) Phylogenetic inference was conducted in MEGA 6 using the maximum likelihood method based on the Kimura 2-parameter. Novel haplotypes identified in this study are in bleu. B) Bacterial DNAs detected in head lice reported in this study and the literature. The pathogenic bacteria in red are those naturally transmitted by body lice to human. 173 / 285 Figure 2. Cytb haplotype networks of human body and head lice. Each circle indicates a unique haplotype and variations in circle size are proportional to haplotype frequencies. Pie colors and sizes in circles represent the continents and the number of their sequence for a haplotype. The length of the links between nodes is proportional to mutational differences. Haplotypes identified in this study are underlined. 174 / 285 Figure 3. Phylogenetic tree highlighting the position of Acinetobacter species identified in head lice from DR Congo. The rpoB sequences were aligned using CLUSTALW, and phylogenetic inferences were conducted in MEGA 6 using the maximum likelihood method based on the Kimura 3-parameter model for nucleotide sequences. Statistical support for internal branches of the trees was evaluated by bootstrapping with 1000 iterations. There were a total of 345 positions in the final dataset. 175 / 285 Table 1. Oligonucleotide sequences of primers and probes used for real-time PCRs and conventional PCRs in this study. Target Name Primers (5’-3’) and probes Source P. humanus F_GATGTAAATAGAGGGTGGTT cytochrome b Duplex A- R_GAAATTCCTGAAAATCAAAC D FAM-CATTCTTGTCTACGTTCATATTTGG-TAMRA VIC-TATTCTTGTCTACGTTCATGTTTGA-TAMRA F_TTAGAGCGMTTRTTTACCC Duplex B- R_AYAAACACACAAAAMCTCCT [7,8] C/E FAM-GAGCTGGATAGTGATAAGGTTTAT-MGB VIC-CTTGCCGTTTATTTTGTTGGGGTTT-TAMRA F_GGTTGGAATTGGATAGTGAT Monoplex R_GGGTCCATAAAGAAATCCG E FAM- TAGGAGGCTTTGTGTGTCTATCCT-TAMRA F_GAGCGACTGTAATTACTAATC Cytb [26] R_CAACAAAATTATCCGGGTCC Acinetobacter spp. F_TACTCATATACCGAAAAGAAACGG [21] rpoB RNA polymerase β R_GGYTTACCAAGRCTATACTCAAC subunit gene FAM-CGCGAAGATATCGGTCTSCAAGC-TAMRA rpoB F_TAYCGYAAAGAYTTGAAAGAAG [28] (zone1) R_CMACACCYTTGTTMCCRTGA R. prowazekii F_AATGCTCTTGCAGCTGGTTCT rOmpB gene R_TCGAGTGCTAATATTTTTGAAGCA [44] ompB FAM-CGGTGGTGTTAATGCTGCGTTACAACA- TAMRA Y. pestis F_ATGGAGCTTATACCGGAAAC plasminogen R_GCGATACTGGCCTGCAAG activator gene PLA FAM-TCCCGAAAGGAGTGCGGGTAATAGG- [44] TAMRA Borrelia spp. F_AGCCTTTAAAGCTTCGCTTGTAG 16S ribosomal Bor16S R_GCCTCCCGTAGGAGTCTGG [45] RNA FAM-CCGGCCTGAGAGGGTGAACGG-TAMRA B. quintana F_TAAACCTCGGGGGAAGCAGA Hypothetical R_TTTCGTCCTCAACCCCATCA yopP intracellular FAM-CGTTGCCGACAAGACGTCCTTG-TAMRA effector [18] 3-oxoacyl-synthase F_GCGGCCTTGCTCTTGATGA gene fabF3 R_GCTACTCTGCGTGCCTTGGA FAM-TGCAGCAGGTGGAGAGAACGTG-TAMRA Anaplasma spp. F_TGACAGCGTACCTTTTGCAT 23S ribosomal TtAna R_TGGAGGACCGAACCTGTTAC [46] RNA FAM-GGATTAGACCCGAAACCAAG-TAMRA C. burnetii F_CAAGAAACGTATCGCTGTGGC Spacers IS1111 R_CACAGAGCCACCGTATGAATC IS1111 [47] FAM-CCGAGTTCGAAACAATGAGGGCTG- TAMRA 176 / 285 Table 2. Number of infested people to single or multiple clades of lice in this study. Clade of lice People infested (n = 27) no. % Single infestation Clade A 8 29.63 Clade D 5 18.52 Clade E 2 7.41 Total 15 55.56 Multiple infestation Clade A/D 12 44.44 Clade A/E 0 0 Clade D/E 0 0 Clade A/D/E 0 0 Total 12 44.44 177 / 285 Table 3. Haplotype frequency of head and body lice identified in 54 head lice. Clade of lice Haplotype Number Acc. no. Clade A A5 16 KM579542 A66 7 MH230928 A67 2 MH230927 A68 2 MH230926 A69 1 MH230925 Clade D D60 3 KX249766 D74 4 MH230924 D75 1 MH230923 D76 8 MH230922 Clade E E62 10 MH230921 Total 54 178 / 285 Table 4. Summary of the bacterial species detected in head lice collected from infested individuals from DR Congo per lice clade. Total Bacterial species Clade of lice (no.) 181 (%) A. baumannii (8) A, (5) D, and (2) E 15 (8.29%) A. pittii (2) A and (1) D 3 (1.66%) A. soli (3) A 3 (1.66%) A. johnsonii (2) A and (1) D 3 (1.66%) A. guillouiae (2) A 2 (1.10%) Candidatus Acinetobacter pediculi (2) A 2 (1.10%) Acinetobacter spp (3) E, (1) A, and (1) D 5 (2.76%) 179 / 285 Chapitre III : Mécanismes de résistance des poux à l’ivermectine 180 / 285 Préambule L’ivermectine est un dérivé semi-synthétique d’une lactone macrocyclique [31]. Son mode d’action est original, car il présente une affinité importante pour les canaux chlorures glutamate-dépendants (GluCl) qui sont présents uniquement chez les invertébrés [31]. Utilisé actuellement dans le traitement de masse de lutte contre les endémies tropicales, efficace contre plusieurs helminthoses [31]. Mais aussi, contre les ectoparasites tels que le sarcopte agent de la gale et les poux, notamment chez les populations de poux ayant développé des résistances envers les insecticides conventionnels [31,32]. Malheureusement, des cas de résistance à l’ivermectine sur le terrain commencent à être signalés [33]. Ainsi, comprendre les mécanismes impliqués dans la résistance à l’ivermectine est une étape clé pour retarder et gérer sa diffusion parmi les populations cibles. Dans un premier temps, nous avons analysé des poux de tête cliniquement résistants à l’ivermectine récoltés sur deux filles du village de Dielmo (Sénégal), traitées à l’ivermectine. Afin d’identifier la présence d’éventuelles mutations sur le GluCl potentiellement associées à la résistance, nous avons cloné puis séquencé le gène en question de ces poux. L’analyse des séquences a permis d’identifier trois mutations faux-sens (A251V, S46P et H272R), uniquement retrouvées chez les poux résistants. Une méthode de PCR-RFLP très sensible et très spécifique a été développée pour la détection de routine et le génotypage des trois mutations sur le terrain. Dans un deuxième temps, nous avons étudié les mécanismes de résistance chez une population de poux de corps de laboratoire chez qui la résistante à l’ivermectine a été sélectionnée par des expositions répétées sur plusieurs générations. Par la suite, sur cette souche sélectionnée, nous avons procédé à la recherche des mutations au niveau de GluCl suivi d’une analyse protéomique globale comparative en comparaison avec la souche sensible de référence. 181 / 285 L’analyse des séquences cDNA de GluCl exprimées chez la souche résistante n’a révélé aucune mutation faux-sens, excluant probablement l'implication de ce gène dans la résistance, au moins dans les conditions de notre expérimentation. L’analyse protéomique comparative entre la souche résistante et la souche sensible a révélé une expression différentielle dans 22 protéines, dont 13 up-régulées et 9 down-régulées chez la souche résistante. Parmi ces protéines, la complexine qui est une protéine de liaison SNARE jouant un rôle clé dans la régulation de la libération des neurotransmetteurs dont le glutamate, a été le plus significativement down-régulée chez la souche résistante comparée à la souche sensible. L’analyse des séquences cDNA des différents variants de complexine exprimés chez la souche résistante a révélé que certains variants présentent une mutation ponctuelle non-sens créant des codons stop et conduisant probablement à la production d’une protéine tronquée, ce qui peut expliquer en partie cette down-régulation. Enfin, nous avons confirmé l'association entre complexine et la résistance à l’ivermectine à travers les ARN interférents et nous avons constaté que la suppression de l’expression de la complexine chez les poux diminue significativement leur sensibilité à l’ivermectine. 182 / 285 Article 9 : Mutations in Ivermectin-target site (GluCl) associated with field- evolved resistance of head lice recovered from Senegal Accepté dans International Journal of Antimicrobial Agents 183 / 285 Mutations in GluCl associated with ivermectin Field-Resistant Head lice from Senegal Nadia Amanzougaghene1, Florence Fenollar2, George Diatta2,3, Cheikh Sokhna2,3, Didier Raoult1, Oleg Mediannikov1* 1Aix-Marseille Univ, IRD, AP-HM, MEPHI, IHU-Méditerranée Infection, Marseille, France 2Aix Marseille Univ, IRD, AP-HM, SSA, VITROME, IHU-Méditerranée Infection, Marseille, France 3VITROME, Campus International UCAD-IRD, Dakar, Senegal *Corresponding author: Dr. Oleg MEDIANNIKOV Phone : +33 (0)4 13 73 24 01, Fax : +33 (0)4 13 73 24 02 and E-mail : [email protected] 184 / 285 Abstract Through its unique mode of action, ivermectin represents a relatively new and very promising tool to fight against human lice, especially in cases of resistance to commonly used pediculicides. However, ivermectin resistance in the field has already begun to be reported. Therefore, understanding the mechanisms involved is a key step in delaying and tackling this phenomenon. In this study, field head lice with confirmed clinical resistance to ivermectin in rural human populations from Senegal were subjected to genetic analysis targeting GluCl gene, the primary target of ivermectin known to be involved in resistance. Through DNA- polymorphism analysis, three relevant non-synonymous mutations in GluCl which were found only in ivermectin-resistant head lice (76 head lice tested), were identified. The A251V mutation found in the TM3 transmembrane domain was the most prevalent (allelic frequency of 0.33), followed by the S46P mutation (0.28) located at the N-terminal extracellular domain. The H272R was in the M3-M4 linker transmembrane region of GluCl and has shown the lowest frequency (0.18). PCR-RFLP diagnostic assays were also developed for their accurate genotyping. Our study is the first to report the presence of GluCl point mutations being detected in clinical ivermectin-resistant head lice occurring in rural human populations of Senegal. Keywords: Head lice; GluCl; Ivermectin; Senegal; Resistance 185 / 285 Introduction The head lice, Pediculus humanus capitis, are obligate blood-sucking parasites that live exclusively in the scalp region of humans [1,2] and represent one of the most prevalent parasitic infestation with a major economic and social concern throughout the world [3]. The recently discovered ability of the head lice to transmit diseases as body lice do, creates a very substantial health risk to infested persons. [4–6] Indeed, such infestations are not controlled in any countries and the outbreaks are still occurring regularly. This is mainly due to the emergence and spread of resistant louse populations to the widely used insecticides which are acetylcholinesterase-inhibiting malathion and synthetic pyrethroid pediculicides, such as permethrin [1,7,8]. This development has prompted research into new treatment strategies such as ivermectin. This drug raised the attention for presenting higher lethal capacity against insects, since it requires only small doses of the product for satisfactory effects [9]. Moreover, ivermectin’s mechanism of action differs from all classes of pediculicides used to treat lice providing opportunities to counter the existing resistance [7,10]. Ivermectin is a macrocyclic lactone, a multifaceted ‘wonder’ drug that had a broad spectrum of activity. It acts robustly against a wide variety of nematode and arthropods [9,11] with its unexpected potential as an antibacterial, antiviral and anti-cancer agent being particularly valuable in improving global public health [9,12,13]. Ivermectin is already deployed and commercialized to treat human lice infestation and several reports indicated that both orally and topically formulations were highly effective in controlling lice infestations [7,14,15]. Ivermectin’s mode of action makes it safer to vertebrates, including humans, as it targets glutamate-gate chloride channel (GluCl) present in invertebrates [11,16]. The channel does not exist in vertebrates and the drug has a low affinity for other vertebrates ligand-gated chloride channels [11]. GluCls contribute extensively to invertebrate nervous system function, including modulating locomotion, regulating feeding and mediating sensory 186 / 285 inputs [17,18]. As a result, modification of channel function by ivermectin action results in paralysis and death [18]. GluCl channels are pentamers belonging to the Cys-loop receptor superfamily, constituted by five subunits organized to form a chloride-permeable pore at the center [19,20]. Each subunit has an extracellular N-terminal domain, which contributes to glutamate-binding site, and four membrane-bound helices (M1–M4) that constitute a channel domain [21,21,22]. The ivermectin-binding site is in the channel domain, lying between M3 and M1 of two adjacent subunits. Ivermectin also makes contact with M2 helice, which lines the pore of ion channel, through its disaccharide moiety and the M2-M3 loop [18]. As with other insecticides, ivermectin is subject to selection pressures that have led to resistance development. Currently, this resistance has been demonstrated in many arthropods and is an increasing problem for their control. Therefore, resolving the mechanisms is an important research priority [18,23,24]. Target-site insensitivity is one of the principal mechanism by which insect pests acquire resistance to insecticides [19]. Indeed, several studies have shown that mutations in GluCl are found to be associated with ivermectin resistance in several arthropods and worms [23–26]. Recently, Diatta et al. (2016) reported a first field-evolved ivermectin-resistance in head lice from Senegal [27]. These lice were recovered from two females after ivermectin treatment. This study aimed to investigate the presence of polymorphisms in the ivermectin- target site GluCl, known to be involved in ivermectin resistance, of these lice. 187 / 285 Materials and methods Study design and louse sampling Samples genotyped in the present study have been collected in November 2015 during an ivermectin oral treatment trial aiming to confirm previous suspicions of ivermectin resistance observed during a first clinical trial conducted in 2014 in the same region, Dielmo a village located in the Sine-Saloum region of Senegal [27]. The full details of the study are described in Diatta et al. (2016) [27]. Briefly, ivermectin (400 µg/kg of body weight) was orally administered in one intake to treat eight females, six with suspected resistance or re- infestation observed during the first trial and two in whom no resistance or re-infestation phenomena were observed. After 24 hours of ivermectin treatment, two cases (2/8; 25%) of clinical ivermectin-resistance (treatment failure) were observed from which six lice alive were recovered. Head lice from all these females (6 lice with clinical resistance and 30 lice dead after ivermectin treatment) and lice collected from the first trial were sent to our laboratory in Marseille to investigate the possible involved mechanism in ivermectin resistance. The Orlando (Culpepper) reference strain of body lice maintained on rabbits, never exposed to ivermectin, was used as a control wild-type. Amplification, cloning and sequencing of GluCl gene Genomic DNA (gDNA) was extracted from louse specimens as described previously [28]. The gDNA sequence of the GluCl gene of P. humanus (GenBank ID: PHUM451790- RA, DS235786) has 7740-bp, consisting of 7 exons and 6 introns. The GluCl fragment (~1722-bp) encompassing all exons, except the first exon which is composed on 3-bps, was PCR amplified using a set of primer pairs listed in table 1. The amplification was performed from a DNA pool of the 6 head lice with suspected ivermectin-resistance and the laboratory susceptible body lice strain to perform DNA polymorphism analysis. 188 / 285 All PCRs were performed using a Peltier PTC-200 model thermal cycler (MJ Research Inc., Watertown, MA, USA) with the Hotstar Taq-polymerase (Qiagen). The purified PCR products were ligated into a pGEMT-easy vector (Promega, Madison, WI) and transformed into JM109 Competent Cells. The plasmid inserts were then PCR amplified using a vector- specific primer set (M13 forward and reverse primers) and subjected to sequencing using the Big Dye Terminator Cycle Sequencing Kit (Perkin Elmer Applied Biosystems, Foster City, CA) with an ABI automated sequencer (Applied Biosystems). The electropherograms were assembled and edited using ChromasPro software (ChromasPro 1.7, Technelysium Pty Ltd., Tewantin, Australia). Analysis of the nucleotide and amino acid sequences was conducted using the ClustalW2 computer program (http://www.ebi.ac.uk/Tools/clustalw2/index.html). Molecular diagnostic assays based on PCR-RFLP for GluCl mutations genotyping Following the identification of mutations potentially associated with ivermectin- resistance, three PCR-RFLP diagnostic assays were designed for their accurate detections. The appropriate restriction enzyme for each mutation was selected using NEBcutter version 2.0 (New England Biolabs, http://nc2.neb.com/NEBcutter2/). Three set pairs of primers for amplifying the fragments encompassing the three mutations were designed using Primer3 software, version 4.0 (http://frodo.wi.mit.edu/primer3/). The details of primers, PCR conditions and restriction enzyme are given in Table 1. The PCR-amplified fragments corresponding to S46P, H272R and A251V polymorphisms were digested with the restriction enzymes AgsI (cut wild type), NsiI (cut wild type) and HhaI (cut mutated type), respectively. Amplicons were subjected to restriction analysis according to the supplier’s instructions. Digested PCR products were separated on 2% agarose gels and the resulting DNA bands were visualized under UV light. In total, 76 head lice with suspected ivermectin-resistance collected from Senegal (six with confirmed clinical resistance and 70 with suspected resistance to ivermectin), including 189 / 285 forty head lice collected from the first ivermectin-trial and thirty-six (including the six with suspected resistance genotyped above) collected from the second trial, were genotyped using these PCR-RFLP essays. Sixteen body lice from the laboratory colony, twenty-four head lice collected in France and Algeria (stored in our laboratory) were also used as possibly susceptible specimens. Statistical analysis The statistically significant difference of the frequency of each mutation between the samples was calculated using a Chi-square test using GraphPad Prism version 7.00 for Windows (GraphPad Software, La Jolla California USA, www.graphpad.com) and P-values of ≤ 0.05 were considered significant. Ethical approval The study protocol was approved by the National Ethics Committee of the Ministry of Health and Social Action in Senegal [no.0267MSAS/DPRS/CNRERS]. Written informed consent was obtained from the individuals involved or from their legal representatives in the case of children. Results Firstly, DNA polymorphism analysis from six-confirmed clinical ivermectin-resistant head lice, recovered from two Senegalese rural girls, and the laboratory susceptible body lice was performed on GluCl gene. Large-scale cloning and sequencing (96 clones from two independent DNA batches) followed by multiple nucleotides and deduced amino acid sequences comparison revealed the presence of five nonsynonymous mutations: serine to proline (S46P), asparagine to aspartic acid (N143D), threonine to alanine (T236A), alanine to valine (A251V) and histidine to arginine (H272R) (numbering based EEB17068; Fig 1B; Fig. S1) resulting from SNPs of thymine to cytosine (T136C), adenine to guanine (A427G), 190 / 285 adenine to guanine (A706G), cytosine to thymine (C752T) and adenine to guanine (A815G), respectively, in coding region of GluCl (numbering based XM00242976; Fig 1A). The N143D and S46P mutations located at the N-terminal extracellular domain of the channel which carries glutamate binding site. The T236A and A251V mutations were found in the TM3 transmembrane domain and the H272R was found in the M3-M4 linker transmembrane region of GluCl. The fact that the N143D mutation occurred in all tested lice, suggests that this mutation is a natural one and has nothing to do with resistance (Table 2). The T236A mutation was found to be located within the binding site of ivermectin, but the fact that it is a conservative amino acid substitution coupled with its low frequency (found only in 3/96 clones) suggests that this mutation is not likely associated with resistance (Fig 1). Therefore, only the three remaining mutations (S46P, A251V and H272R), found from the head lice with clinical ivermectin-resistance, were thought to be potentially associated with ivermectin-resistance. Secondly, to investigate the occurrence of these three mutations (S46P, A251V and H272R) in all louse specimens collected during the two ivermectin clinical trials from Senegal (76 lice) and in order to support their association with ivermectin resistance, three RFLP-PCR essays for genotyping were developed. First, these RFLP-PCR essays were approved by testing the known genotype clones. The results have shown that the genotypes of three mutations can be detected accurately by these essays (Fig 2). RFLP-PCR essays were then used to genotype louse individuals from the studied louse specimens. Louse specimens never exposed to ivermectin were used as susceptible controls (40 lice). As shown in table 3, none of the three mutations were found in the laboratory susceptible colony (8 body lice tested) or in the other louse specimens used as susceptible controls (32 lice tested), while, all the three mutations were detected in clinical ivermectin- resistant head lice collected in Senegal during the two ivermectin-clinical trials (76 head lice 191 / 285 analyzed). The most prevalent mutation was A251V with allelic frequency of 0.33 followed by S46P (0.28) and H272R (0.18) mutations. Considering the years of sampling, the allelic frequency of A251V mutation was significantly elevated from 2014 to 2015, the opposite scenario was observed for H272R mutation, while for S46P the allelic frequency remained stable over the two years. Discussion and conclusion In this study, a field head louse with confirmed clinical resistance to ivermectin in rural human populations of Senegal [27] were subjected to genetic analysis targeting GluCl gene, the primary target of ivermectin, known to be involved in resistance in many arthropods and nematode [21,23,25,26,29]. Through DNA-polymorphism analysis (cloning and sequencing combined with PCR-RFLP genotyping), three relevant non-synonymous mutations in GluCl gene, found only in clinical ivermectin-resistant head lice, were identified. The A251V mutation found in the TM3 transmembrane domain was the most prevalent, followed by the S46P mutation located at the N-terminal extracellular domain. The H272R was in the M3-M4 linker transmembrane region of GluCl and showed the low frequency. Overall, all mutation frequencies were low possibly suggesting that a selection could be acting on this gene as a consequence of ongoing resistance. Several studies on GluCls have identified different point mutations affecting ivermectin sensitivity in arthropods [21]. In Drosophila melanogaster, P299S mutation located in the M2-M3 linker region of GluCl, has been associated with ivermectin resistance [25], while the A309V mutation in the TM3 transmembrane domain of Plutella xylostella GluCl, has been associated with a 10-fold reduction in abamectin sensitivity [24]. In spotted spider mite, Tetranychus urticae, the TuGluCl1 G323D and TuGluCl3 G326E mutations found in the 192 / 285 TM3 transmembrane domain, are shown to be associated with 18- and 2000-fold resistance to abamectin, respectively [21,23]. The three mutations we reported herein do not map onto any of the previously reported mutations in other arthropods. Furthermore, whether these mutations contribute directly to ligand binding, or whether they cause conformational change that influence ligand binding, is currently not known, and requires further characterization. The most prevalent mutation A251V is located at TM3, which is important in the chloride channel pore formation [19,20], and more importantly, it is near critical residue that drastically affect ivermectin potency, which is TM3-Gly36’ residue (corresponding to the Gly232 in P. humanus GluCl) identified by site-directed mutagenesis in the avr-14b subunit of Haemonchus contortus [30]. The mutation in this residue was also associated with elevated level of abamectin resistance in wild isolates T. urticae [21,23]. Even though this residue occurs at the ivermectin binding site, the mutation in this site was found to reduce ivermectin sensitivity by altering the functional properties of the GluCl rather than specifically affecting the binding of ivermectin [31]. Similar mechanism can be suggested to the A251V mutation where alanine replaced by valine, which is likely to have significant biophysical and thus structural consequences for the P. humanus GluCl with resulting implications for the efficacy of ivermectin binding. Although the mutation S46P is located in the glutamate binding site at the N-terminal extracellular domain of GluCl and not in or near reported putative ivermectin binding sites, we cannot exclude the possibility that it may be involved in the resistance because N- terminus region can be a structural requirement for ivermectin-induced activation of the GluCl (33–35). Moreover, in the Cooperia onchophora nematode, three (E114G, V235A and L256F) and two (V60A and R101H) point mutations were found at the N-terminal extracellular domain of GluCla3 and GluClb, respectively [26]. Amongst these mutations, 193 / 285 electrophysiological studies determined that the L256F mutation of GluCla3 causes a 3-fold reduction in ivermectin sensitivity [26]. The glutamate binding site is also site that harbour mutations in wild avermectins resistant strains of nematode Caenorhabditis elegans [29]. In summary, our study has for the first time associated the possible involvement of three-point mutations on GluCl gene in clinical ivermectin-resistant head lice occurring in rural human populations of Senegal. No similar mutations have been reported in other studies so far. Therefore, functional expression and analysis of GluCl variants containing these mutations would provide crucial information on the toxicodynamic role of these mutations in determining ivermectin sensitivity. 194 / 285 Acknowledgments The authors thank the populations of Dielmo for allowing them to conduct the clinical trial. Funding: This study was supported by IHU-Méditerranée Infection and the National Research Agency under the program « Investissements d’avenir », reference ANR-10-IAHU- 03. 195 / 285 References [1] Bonilla DL, Durden LA, Eremeeva ME, Dasch GA. The biology and taxonomy of head and body lice--implications for louse-borne disease prevention. PLoS Pathog 2013;9:e1003724. doi:10.1371/journal.ppat.1003724. [2] Veracx A, Raoult D. Biology and genetics of human head and body lice. Trends Parasitol 2012;28:563–71. doi:10.1016/j.pt.2012.09.003. [3] Chosidow O. Scabies and pediculosis. The Lancet 2000;355:819–26. doi:10.1016/S0140-6736(99)09458-1. [4] Raoult D, Abat C. Developing new insecticides to prevent chaos: the real future threat. Lancet Infect Dis 2017;17:804–5. doi:10.1016/S1473-3099(17)30395-X. [5] Amanzougaghene N, Fenollar F, Sangaré AK, Sissoko MS, Doumbo OK, Raoult D, et al. Detection of bacterial pathogens including potential new species in human head lice from Mali. PLOS ONE 2017;12:e0184621. doi:10.1371/journal.pone.0184621. [6] Angelakis E, Diatta G, Abdissa A, Trape J-F, Mediannikov O, Richet H, et al. Altitude- dependent Bartonella quintana genotype C in head lice, Ethiopia. Emerg Infect Dis 2011;17:2357–9. doi:10.3201/eid1712.110453. [7] Chosidow O, Giraudeau B, Cottrell J, Izri A, Hofmann R, Mann SG, et al. Oral ivermectin versus malathion lotion for difficult-to-treat head lice. N Engl J Med 2010;362:896–905. doi:10.1056/NEJMoa0905471. [8] Durand R, Bouvresse S, Berdjane Z, Izri A, Chosidow O, Clark JM. Insecticide resistance in head lice: clinical, parasitological and genetic aspects. Clin Microbiol Infect Off Publ Eur Soc Clin Microbiol Infect Dis 2012;18:338–44. doi:10.1111/j.1469- 0691.2012.03806.x. [9] Crump A. Ivermectin: enigmatic multifaceted ‘wonder’ drug continues to surprise and exceed expectations. J Antibiot (Tokyo) 2017;70:495–505. doi:10.1038/ja.2017.11. 196 / 285 [10] Yoon KS, Strycharz JP, Baek JH, Sun W, Kim JH, Kang JS, et al. Brief exposures of human body lice to sub-lethal amounts of ivermectin over transcribes detoxification genes involved in tolerance. Insect Mol Biol 2011;20:687–99. doi:10.1111/j.1365- 2583.2011.01097.x. [11] Laing R, Gillan V, Devaney E. Ivermectin - Old Drug, New Tricks? Trends Parasitol 2017;33:463–72. doi:10.1016/j.pt.2017.02.004. [12] Pettengill MA, Lam VW, Ollawa I, Marques-da-Silva C, Ojcius DM. Ivermectin inhibits growth of Chlamydia trachomatis in epithelial cells. PloS One 2012;7:e48456. doi:10.1371/journal.pone.0048456. [13] Mastrangelo E, Pezzullo M, De Burghgraeve T, Kaptein S, Pastorino B, Dallmeier K, et al. Ivermectin is a potent inhibitor of flavivirus replication specifically targeting NS3 helicase activity: new prospects for an old drug. J Antimicrob Chemother 2012;67:1884–94. doi:10.1093/jac/dks147. [14] Foucault C, Ranque S, Badiaga S, Rovery C, Raoult D, Brouqui P. Oral ivermectin in the treatment of body lice. J Infect Dis 2006;193:474–6. doi:10.1086/499279. [15] Pariser DM, Meinking TL, Bell M, Ryan WG. Topical 0.5% ivermectin lotion for treatment of head lice. N Engl J Med 2012;367:1687–93. doi:10.1056/NEJMoa1200107. [16] Chaccour C, Hammann F, Rabinovich NR. Ivermectin to reduce malaria transmission I. Pharmacokinetic and pharmacodynamic considerations regarding efficacy and safety. Malar J 2017;16:161. doi:10.1186/s12936-017-1801-4. [17] Wolstenholme AJ. Glutamate-gated Chloride Channels. J Biol Chem 2012;287:40232– 8. doi:10.1074/jbc.R112.406280. [18] Wang X, Puinean AM, O´Reilly AO, Williamson MS, Smelt CLC, Millar NS, et al. Mutations on M3 helix of Plutella xylostella glutamate-gated chloride channel confer 197 / 285 unequal resistance to abamectin by two different mechanisms. Insect Biochem Mol Biol 2017;86:50–7. doi:10.1016/j.ibmb.2017.05.006. [19] Ozoe Y. Chapter Four - γ-Aminobutyrate- and Glutamate-gated Chloride Channels as Targets of Insecticides. In: Cohen E, editor. Adv. Insect Physiol., vol. 44, Academic Press; 2013, p. 211–86. doi:10.1016/B978-0-12-394389-7.00004-1. [20] Lynagh T, Webb TI, Dixon CL, Cromer BA, Lynch JW. Molecular Determinants of Ivermectin Sensitivity at the Glycine Receptor Chloride Channel. J Biol Chem 2011;286:43913–24. doi:10.1074/jbc.M111.262634. [21] Dermauw W, Ilias A, Riga M, Tsagkarakou A, Grbić M, Tirry L, et al. The cys-loop ligand-gated ion channel gene family of Tetranychus urticae: implications for acaricide toxicology and a novel mutation associated with abamectin resistance. Insect Biochem Mol Biol 2012;42:455–65. doi:10.1016/j.ibmb.2012.03.002. [22] Fuse T, Kita T, Nakata Y, Ozoe F, Ozoe Y. Electrophysiological characterization of ivermectin triple actions on Musca chloride channels gated by l-glutamic acid and γ- aminobutyric acid. Insect Biochem Mol Biol 2016;77:78–86. doi:10.1016/j.ibmb.2016.08.005. [23] Kwon DH, Yoon KS, Clark JM, Lee SH. A point mutation in a glutamate-gated chloride channel confers abamectin resistance in the two-spotted spider mite, Tetranychus urticae Koch. Insect Mol Biol 2010;19:583–91. doi:10.1111/j.1365-2583.2010.01017.x. [24] Wang X, Wang R, Yang Y, Wu S, O’Reilly AO, Wu Y. A point mutation in the glutamate-gated chloride channel of Plutella xylostella is associated with resistance to abamectin. Insect Mol Biol 2016;25:116–25. doi:10.1111/imb.12204. [25] Kane NS, Hirschberg B, Qian S, Hunt D, Thomas B, Brochu R, et al. Drug-resistant Drosophila indicate glutamate-gated chloride channels are targets for the antiparasitics 198 / 285 nodulisporic acid and ivermectin. Proc Natl Acad Sci U S A 2000;97:13949–54. doi:10.1073/pnas.240464697. [26] Njue AI, Hayashi J, Kinne L, Feng X-P, Prichard RK. Mutations in the extracellular domains of glutamate-gated chloride channel alpha3 and beta subunits from ivermectin- resistant Cooperia oncophora affect agonist sensitivity. J Neurochem 2004;89:1137–47. doi:10.1111/j.1471-4159.2004.02379.x. [27] Diatta G, Abat C, Sokhna C, Tissot-Dupont H, Rolain J-M, Raoult D. Head lice probably resistant to ivermectin recovered from two rural girls in Dielmo, a village in Sine-Saloum, Senegal. Int J Antimicrob Agents 2016;47:501–2. doi:10.1016/j.ijantimicag.2016.03.013. [28] Amanzougaghene N, Akiana J, Mongo Ndombe G, Davoust B, Nsana NS, Parra H-J, et al. Head Lice of Pygmies Reveal the Presence of Relapsing Fever Borreliae in the Republic of Congo. PLoS Negl Trop Dis 2016;10:e0005142. doi:10.1371/journal.pntd.0005142. [29] Ghosh R, Andersen EC, Shapiro JA, Gerke JP, Kruglyak L. Natural variation in a chloride channel subunit confers avermectin resistance in C. elegans. Science 2012;335:574–8. doi:10.1126/science.1214318. [30] Lynagh T, Lynch JW. Ivermectin binding sites in human and invertebrate Cys-loop receptors. Trends Pharmacol Sci 2012;33:432–41. doi:10.1016/j.tips.2012.05.002. [31] Atif M, Estrada-Mondragon A, Nguyen B, Lynch JW, Keramidas A. Effects of glutamate and ivermectin on single glutamate-gated chloride channels of the parasitic nematode H. contortus. PLOS Pathog 2017;13:e1006663. doi:10.1371/journal.ppat.1006663. 199 / 285 Fig 1. Mutations in GluCl associated with ivermectin field-resistant head lice in rural human populations of Senegal. (A) Map of collection site of ivermectin-resistant head lice in rural human populations of Dielmo village located in the Sine-Saloum, Senegal. (B) Genetic structure of the GluCl gene and (C) schematic diagram of the predicted structure of the GluCl protein showing the position of five mutations. Positions of mutations are indicated by stars. Amino acid replacements that may result in structural alterations and found only in head lice with clinical ivermectin-resistance are indicated by red. Amino acid positions are numbered according to body lice GluCl protein (accession no. EEB17068). 200 / 285 Fig 2. PCR-RFLP essays for genotyping S46P, H272R and A251V mutations of the GluCl gene. A) (Upper) The AgsI, NsiI and HhaI recognition sites are shown above the DNA sequences. (Lower) The size of the expected fragments for all possible genotypes. B-D) SYBR safe- stained 2% agarose gels showing the results of digestion with AgsI, NsiI and HhaI enzymes. 201 / 285 Table 1. Primer sequences used in this study. Purpose Primer Primer sequences (5′-3′) Product size (bp) name GluCl 6079F CAATTAATTCGACGGATTCAG 1722 7800R GATTGATTTACCAACGACGGC Sequencing 7157F AGTGACAACATTACTCACAA of 1722-bp GluCl 6574F GCTCACTTCGAATGGCCAGTTG 6686R CCATCCCTACGACCATCAAATT 7287R CTAACAAAGCTCCAAATACGAAC PCR-RFLP 5794F AACCAAACGGATAAAACCGATG 865 (for S46P genotyping) (AgsI: 550+315) * 6658R CCGACCAAAAGCTTTAAATTAT 6833F AAGTTGGCGATCCCGTTCAAG 919 (for H272R genotyping) (NsiI: 379+540) * 7750R CAATCGAATTAATTATCTTCCGT 7203F GCATCATTGCCACCGGTA 547 (for A251V genotyping) (HhaI: 440+107) * 7750R CAATCGAATTAATTATCTTCCGT * Length of fragment obtained after digestion with the corresponding restriction enzyme endonuclease 202 / 285 Table 2. The five nonsynonymous mutations identified in GluCl gene. GluCl Nucleotide Amino acid Localization of Louse population Pattern of mutation mutation change mutation (ivermectin-HL or substitutions Lab-BL) Mut-1 T136C S46P N-terminal ivermectin-HL Transition extracellular domain Mut-2 A427G N143D N-terminal ivermectin-HL / Lab- Transition extracellular domain BL Mut-3 A706G T236A M3 transmembrane ivermectin-HL Transition domain Mut-4 C752T A251V M3 transmembrane ivermectin-HL Transition domain Mut-5 A815G H272R M3-M4 linker ivermectin-HL Transition HL: head lice; BL: body lice 203 / 285 Table 3. Frequencies of S46P, A251V and H272R mutations using the RFLP-PCR essays in louse specimens. Lice tested (no.) Genotype Genotype frequencies: no. (%) profile S46P A251V H272R HL-Senegal* RR 3 (8.3%) 6 (16.6%) 0 (0%) (ivermectin-trial RS 16 (44.4%) 20 (55.5%) 6 (16.7%) 2015) (36) SS 17 (47.2%) 10 (27.8%) 30 (83.3%) Mutated allelic frequency 0.30 - a 0.44§↑b 0.08§↓c HL-Senegal* RR 0 (0%) 3 (7.5%) 2 (5%) (ivermectin-trial RS 21 (52.5%) 12 (30%) 17 (42.5%) 2014) (40) SS 19 (47.5%) 25 (62.5%) 21 (52.5%) Mutated allelic frequency 0.26 a 0.22 b 0.26 c RR 3 (3.9%) 9 (11.8%) 2 (2.6%) Total* (76) RS 37 (48.7%) 32 (42.1%) 23 (30.3%) SS 36 (47.4%) 35 (46.1%) 51 (67.1%) Mutated allelic frequency 0.28 0.33 0.18 RR 0 (0%) 0 (0%) 0 (0%) HL-Algeria (16) RS 0 (0%) 0 (0%) 0 (0%) SS 16 (100%) 16 (100%) 16 (100%) Mutated allelic frequency 0 0 0 RR 0 (0%) 0 (0%) 0 (0%) HL-France (8) RS 0 (0%) 0 (0%) 0 (0%) SS 8 (100%) 8 (100%) 8 (100%) Mutated allelic frequency 0 0 0 RR 0 (0%) 0 (0%) 0 (0%) BL-SDF (8) RS 0 (0%) 0 (0%) 0 (0%) SS 8 (100%) 8 (100%) 8 (100%) Mutated allelic frequency 0 0 0 RR 0 (0%) 0 (0%) 0 (0%) BL-Laboratory RS 0 (0%) 0 (0%) 0 (0%) colony (8) SS 8 (100%) 8 (100%) 8 (100%) Allelic frequency 0 0 0 *Lice specimens with suspected IVERMECTIN -resistance; HL: head lice; BL: body lice. RR, resistant homozygote in which both alleles are mutated; SS, susceptible homozygote in which neither allele is mutated; RS, heterozygote in which only one of the two alleles is mutated. A Chi-2 test comparison of the allele frequencies was carried out between HL-Senegal collected in 2014 and 2015. Significant levels: §↑bp < 0.05 compared with b, §↓c p < 0.05 compared with c. 204 / 285 Supporting Information Ph-GluCl ------Dm-GluCl MGSGHYFWAILYFASLCSASLANNAKVNFREKEKKVLDQILGAGKYDARIRPSGINGTDG Ph-GluCl ------MEYSVQLTFREQWLDERLKFNDFEGRIKYLTLTDANRVWM Dm-GluCl PAIVRINLFVRSIMTISDIKMEYSVQLTFREQWTDERLKFDDIQGRLKYLTLTEANRVWM ************* ******:*::**:******:****** Ph-GluCl PDLFFSNEKEGHFHNIIMPNVYIRIFPHGSVLYSIRISLTLSCPMNLKLYPLDRQVCSLR Dm-GluCl PDLFFSNEKEGHFHNIIMPNVYIRIFPNGSVLYSIRISLTLACPMNLKLYPLDRQICSLR ***************************:*************:*************:**** Ph-GluCl MASYGWTTADLVFLWKVGDPVQVVKNLHLPRFTLEKFFTDYCNSKTNTGEYSCLKVDLLF Dm-GluCl MASYGWTTNDLVFLWKEGDPVQVVKNLHLPRFTLEKFLTDYCNSKTNTGEYSCLKVDLLF ******** ******* ********************:********************** TM1 TM2 Ph-GluCl KREFSYYLIQIYIPCCMLVIVSWVSFWLDQSAVPARVSLGVTTLLTMATQTSGINASLPP Dm-GluCl RREFSYYLIQIYIPCCMLVIVSWVSFWLDQGAVPARVSLGVTTLLTMATQTSGINASLPP :***************************** ***************************** TM3 Ph-GluCl VSYTKAIDVWTGVCLTFVFGALLEFALVNYASRS-----DMHRENMKKQRRQCELEHAAS Dm-GluCl VSYTKAIDVWTGVCLTFVFGALLEFALVNYASRSGSNKANMHKENMKKKRRDLE---QAS ********************************** :**:*****:**: * ** Ph-GluCl LEAAADLLE-DGATTFAMKPLVGHRGDALAIEKARQCEIHMQ-PKRDDCCRTWISKFPT- Dm-GluCl LDAASDLLDTDSNATFAMKPLVRHPGDPLALEKRLQCEVHMQAPKRPNCCKTWLSKFPTR *:**:***: *. :******** * **.**:** ***:*** *** :**:**:***** TM4 Ph-GluCl ---RSKRIDVISRITFPLVFALFNVVYWSTYLFREDTEDN Dm-GluCl QCSRSKRIDVISRITFPLVFALFNLVYWSTYLFREEEDE- *********************:**********: :: Figure S1. Amino acid sequences alignment of P. humanus and D. melanogaster GluCls. The five mutations are indicated by a triangle. The three mutations in red are those that may result in structural alterations and found only in head lice with suspected IVERMECTIN - resistance. The delineation of the four membrane-bound helices (TM1-TM4) is based on the crystal structure of the Caenorhabditis elegans GluClα (32). Ph: P. humanus (accession no. EEB17068); Dm: D. melanogaster (accession no. AAC47266). 205 / 285 Article 10 : Complexin in Ivermectin resistance in body lice Accepté avec révisions mineures dans PLOS Genetics 206 / 285 Complexin in ivermectin resistance in body lice Nadia Amanzougaghene1, Florence Fenollar2, Claude Nappez2, Amira Ben-Amara1, Philippe Decloquement1, Said Azza1, Yassina Bechah2, Eric Chabrière1, Didier Raoult1*, Oleg Mediannikov1* 1Aix Marseille Univ, IRD, APHM, MEPHI, IHU-Méditerranée Infection, Marseille, France 2Aix Marseille Univ, IRD, APHM, VITROME, IHU-Méditerranée Infection, Marseille, France *Corresponding authors: Dr. Oleg MEDIANNIKOV Address: MEPHI, IRD, APHM, IHU-Méditerranée Infection, 19-21 Boulevard Jean Moulin, 13385 Marseille Cedex 05 Tel: +33 (0)4 13 73 24 01, Fax: +33 (0)4 13 73 24 02, E-mail: [email protected], Prof. Didier RAOULT Address: MEPHI, IRD, APHM, IHU-Méditerranée Infection, 19-21 Boulevard Jean Moulin, 13385 Marseille Cedex 05 Tel: +33 (0)4 13 73 24 01, Fax: +33 (0)4 13 73 24 02, E-mail: [email protected] Word abstract count: 247 Word text count: 4,461 Keywords: Body lice, Ivermectin, Resistance, proteomic, Complexin, RNAi 207 / 285 Abstract Ivermectin (IVM) has emerged as very promising pediculicide, particularly in cases of resistance to commonly used pediculicides. Recently, however, the first field-evolved IVM- resistance in lice was reported. To gain insight into the mechanisms underlying IVM- resistance, we both looked for mutations in the IVM-target site (GluCl) and searched the entire proteome for potential new loci involved in resistance from laboratory susceptible and IVM-selected resistant body lice. Polymorphism analysis of cDNA GluCl showed no non-silent mutations, most likely excluding its involvement in IVM-resistance. Proteomic analysis identified 22 differentially regulated proteins, of which 13 were upregulated and 9 were downregulated in the resistant strain. We evaluated the correlation between mRNA and protein levels by qRT-PCR and found that the trend in transcriptional variation was consistent with the proteomic changes. Among differentially expressed proteins, a complexin (Cpx) i.e. a neuronal protein which plays a key role in regulating neurotransmitter release, was shown to be the most significantly down-expressed in the IVM-resistant lice. Moreover, DNA-mutation analysis revealed that some Cpx transcripts from resistant lice gained a premature stop codon, suggesting that this down-expression might be due, in part, to secondary effects of a nonsense mutation inside the gene. We further confirmed the association between Cpx and IVM-resistance by RNA- interfering and found that knocking down the Cpx expression induces resistance to IVM in susceptible lice. Our results provide evidence that Cpx plays a significant role in regulating IVM resistance in body lice and represents the first evidence that links Cpx to insecticide resistance. 208 / 285 Introduction Sucking lice (Anoplura) are obligate blood-feeding ectoparasites of eutherian mammals [1]. Humans are the preferred host for two species: Phthirus pubis and Pediculus humanus [2,3]. The latter has significant relevance to public health and includes two ecotypes: head lice (P. h. capitis), which live in the hair, and body lice (P. h. humanus), which live in clothing [1,3,4]. Head lice are common and can be found worldwide [1], with children being at increased risk [2]. Conversely, body lice are associated with poor socio-economic conditions [1,4] and homeless people and refugee-camp populations are predominantly affected [4,5]. Body lice are the main vectors of at least three dangerous pathogenic bacteria, namely: Rickettsia prowazekii, Bartonella quintana and Borrelia recurrentis [1,4]. The prevalence of the body louse is underestimated in many developed countries and, as the number of homeless people increases, louse-borne infectious diseases are also on the rise [1,5]. Recent studies showed that head lice also have the ability to transmit bacterial diseases [6,7]. This poses a very substantial threat to humanity, because such infestations are not controlled in any countries, including developed nations, despite repeated efforts to eradicate them [7]. This is mainly due to the resistance developed by lice to widely-used insecticides such as malathion and pyrethroid [1,8]. The use of new effective products with different modes of action, such as ivermectin (IVM), have proven to be the most promising alternative to combating the problem of resistance [1,9] IVM belongs to the macrocyclic lactone complex [10,11] and blocks synaptic transmission in invertebrates by binding to glutamate-gated chlorine channels (GluCls) in nerves and muscles, which are its primary target, leading to hyperpolarization, paralysis and death [10]. GluCls are not present in vertebrates and, as such, are thought to confer the broad safety margin of IVM [11]. IVM was the world’s first endectocide, capable of killing a wide 209 / 285 variety of parasites and vectors, including lice [12]. IVM is already used to treat human lice and several reports indicated that both orally and topically administered formulations were highly effective in controlling active both body and head lice infestations [9,13–15]. Currently, resistance to IVM has been widely demonstrated in many arthropods and is an increasing problem for their control [16,17]. Recently, field evolved resistance to IVM in head lice was documented in Senegal, for the first time, and was reported to cause reduced field control efficacy [18]. Understanding the mechanisms of IVM resistance is, therefore, a key step in delaying and tackling this phenomenon. IVM resistance in arthropods has been associated with several mechanisms, including reduced cuticular penetration [16], mutation in the target site [19] and metabolic resistance due to the overexpression of xenobiotic pumps from the ABC family [16,17,20]. Although mechanisms of IVM-resistance in lice have not yet been elucidated, in an attempt to identify inducible metabolic factors involved in IVM- tolerance, Yoon et al., showed that IVM induced detoxification genes, including ATP binding cassette and cytochrome P450, suggesting their association with its xenobiotic metabolism, thereby resulting in tolerance [21]. To gain a deeper understanding of mechanisms underlying IVM resistance in lice, we analyzed the IVM-target site from laboratory susceptible and IVM-selected resistant strains. Additionally, we used functional proteomics and performed a global proteomic analysis between the two strains. In addition, we assessed the correlation between mRNA and protein levels for differentially expressed proteins using quantitative real-time PCR, and further verified the functionality of a key candidate gene by RNA interference (RNAi). Results 1. Resistance levels of selected laboratory lice against IVM From the body lice susceptible strain (Orlando strain; Lab-IVS), IVM resistant selection (Lab-IVR) was successfully achieved by continuous exposure to IVM for ten 210 / 285 generations in the laboratory. The LT50 value for Lab-IVS strain was 28.83 hours (24.47- 32.78 hours), and for the Lab-IVR strain, 157 hours (144.91-172.37) (Table 1). The Lab-IVR strain exhibited 5.4-fold greater resistance against IVM when compared with the reference Lab-IVS strain, suggesting that the Lab-IVR strain had developed low and moderate resistance. 2. Cloning the body louse cDNA GluCl and polymorphism analysis The open reading frame (ORF) of the body louse GluCl was composed of 1,110 nucleotides encoding 369 amino acids. Analysis of the polymorphism patterns of the cDNA sequences from the Lab-IVR and Lab-IVS strains showed the presence of six-point mutations (the Lab-IVR strain showed all mutations and the Lab-IVS carried only two mutations), all were silent substitutions. 3. Identification of differentially expressed proteins between IVM-resistant and susceptible lice Differentially expressed proteins from the Lab-IVS and Lab-IVR strains were identified and quantified by label-free Nano-LC-MS/MS analysis. In total, 407 proteins were identified, including 22 which were differentially expressed, of which 13 were up-regulated and 9 were down-regulated in the resistant Lab-IVR strain (Table 2). Gene ontology annotation, including molecular function, biological process and cellular component, was conducted to categorize these proteins (Table 3 and Fig S1). The main molecular functions were catalytic activity and binding for both down- and up-regulated proteins. The cellular components of down- and up-regulated proteins were mainly cell, membrane, organelle and macromolecular complex. According on biological process, the proteins were mainly classified in cellular process, single-organism process and metabolic process (Fig S1 and Fig S2). 4. Validation of proteomics data at the RNA level by qRT-PCR 211 / 285 To evaluate the proteomic data and the correlation between mRNA and protein levels, we selected 15 differentially expressed genes to quantify their relative expression levels by qRT-PCR. As shown in Fig 1, the trend in transcriptional variation for all the selected proteins was consistent with the proteomic changes determined in the proteomic analysis, suggesting that this method is a reliable way of identifying and quantifying differentially expressed proteins in lice. Of all the differentially expressed proteins, complexin (Cpx) showed the most dramatically altered expression at proteomic level (10-fold down-regulated in Lab-IVR strain), which was correlated at the mRNA level with a slight difference (3.4-fold down- regulated in Lab-IVR strain). Furthermore, because of its impact on neuronal functions as the key regulators of neurotransmitter release [22] we suggest that this gene may play a significant role in regulating the IVM resistance mechanism. Thus, a Cpx was selected as our candidate gene for the subsequent functional verification. Fig 1. Comparison of proteomic and qRT-PCR results. The x-axis shows the 15 selected genes, while the y-axis gives the fold change observed for the Lab-IVR vs the Lab- IVS strains. EF1α was used to normalize the mRNA levels. Values are means ±SEMs (n=3). The selected genes are: Cpx: Complexin, Tryp: Trypsin, CLTC: Clathrin heavy chain, RPS3A: 40S ribosomal protein S3a, TUBα1: Tubulinα1, OAT: Ornithine aminotransferase, HSP: Heat shock protein, Idh: Isocitrate dehydrogenase subunit, Adk: Adenylate kinase, ATPase: ATP synthase, Fib-H: Heavy-chain filboin, MO-porin: Mitochondrial outer membrane porin channel, Lmpt: Limpet, Tubß2: Tubulinß2 and E0W0W7. 5. Characterization of complete Cpx cDNA and its relationship to other Cpxs 212 / 285 The complete cDNA sequence of the Cpx was obtained by RT-PCR and RACE methods based on the partial sequence annotated from the body lice genome sequencing project. Its cDNA contains 426-bps open reading frame encoding 141 amino acids residues. A multiple sequence alignment of lice Cpx with insect and worms Cpxs demonstrates that homology is particularly high in the central predicted SNARE-binding domain (boxed) and in adjacent regions (Fig 2B). Homology analysis of amino acid sequence indicated that lice Cpx shared 91.5% identity with the ortholog in Cyphomyrmex costatus, 87.9% identity with Anoplophora glabripennis, 73.4% identity with Drosophila melanogaster and 44.7% identity with Caenorhabditis elegans. Phylogenetic relationships showed that lice Cpx clustered with insect Cpx and had the highest homology with C. costatus Cpx (Fig 2A). Fig 2. Comparison of Cpx proteins. (A) Phylogenetic tree showing the phylogenetic relationships of Cpx genes from Insecta and non-Insecta species. The tree was generated by ClustalW alignment of the amino acid sequences of Cpx genes using the neighbor-joining (NJ) method. (B) Amino acid sequence alignment of P. humanus (Ph) with members of the Cpx family. Identical residues are marked green and highly conserved residues are marked red. The blue box indicates the position of the predicted SNARE-binding domain. Dm: Drosophila melanogaster; Ce: Caenorhabditis elegans, Cc: Cyphomyrmex costatus, Ag: Anoplophora glabripennis. 6. Cloning and polymorphism analysis of Cpx transcripts To investigate whether mutations in the Cpx gene are involved in the mRNA downregulation leading to a decrease in protein expression observed in Lab-IVR, full-length ORF Cpx cDNA sequences were compared between the Lab-IVR and Lab-IVS strains. Large-scale cloning and sequencing (48 clones from two independent cDNA batches) 213 / 285 followed by multiple sequence comparison revealed the presence of a one nucleotide base pair insertion (A) at position 292-bps. The insertion was found on 17 clones out of 48 analyzed only from the Lab-IVR. This insertion causes a frameshift starting at amino acid 100 within the C-terminal domain of Cpx and results in a premature stop codon at amino acid 111 compared to normal Cpx (Fig S3). 7. Knockdown of Cpx gene by RNAi and subsequent decrease in the sensitivity of lice to IVM To further investigate the function of Cpx, we used RNAi technology to knockdown the expression of this gene in the susceptible Lab-IVS strain. The results showed that the Cpx mRNA levels reduced significantly, by 16.34% at 24 hours post-injection of dsRNA-Cpx and reached maximal reduction by 75.52% at 48 hours post-injection compared to the control injected with dsRNA-plasmid (pQE-30) (Fig 3A), indicating that this gene was mostly silenced by RNAi. No apparent physiological alterations were noticed in lice injected either with dsRNA Cpx or pQE-30 compared to the control non-injected lice. These findings demonstrate that dsRNA injection-based RNAi resulted in the knockdown of the Cpx gene in body lice. To determine whether knocking down the expression of Cpx decrease the susceptibility to IVM in the susceptible Lab-IVS strain, we performed bioassays to compare IVM resistance levels among Lab-IVS lice at 48 hours post-injection with either dsRNA Cpx or pQE-30. The results showed that the mortality rates in lice down-expressing Cpx were significantly reduced compared to the control from 24 hours post exposure to a lethal dose of IVM (150 µg/kg) (P < 0.05, Fig 3B). Based on LT50, the IVM susceptibility of dsRNA-Cpx lice decreased by 3.2-fold (LT50 = 92.54 hours; Table 4) compared to the reference susceptible Lab-IVS (LT50 = 28.8 hours; Table 1) and it was only 2.2-fold higher than that in the Lab-IVR strain (LT50 = 157 hours, RR= 5.4; Table 1). Taken together, the RNAi- 214 / 285 mediated knockdown of Cpx decreased the lice’s susceptibility to IVM (increased their resistance), providing relatively convincing evidence that this gene contributes to IVM resistance in this resistant strain. Fig 3. The effect of dsRNA on expression of Cpx in the IVM-susceptible strain and the effect of Cpx knockdown on IVM resistance. (A) Cpx mRNA levels were quantified by qRT-PCR in 72 hours of injection of dsRNA Cpx or pQE30 (control). The change in mRNA levels in the ds-RNA Cpx were calculated relative to controls. Values are means +SEMs (n=3). Asterisks (*) indicate that Cpx dsRNA significantly suppresses the levels of Cpx transcript (t-test, *P<0.05 and **P<0.01). (B) Bioassays for Lab-IVS lice exposed to IVM (150 µg/Kg) started 48 hours post-injection of dsRNA Cpx or pQE30 (control). Asterisks (*) indicate that knocking down Cpx expression decreased IVM-susceptibility in the Lab-IVS strain compared to the control (Chi-2 test, *P<0.05 and **P<0.01). Discussion and conclusions IVM is a very promising tool to fight different infestations such as, for instance, pediculosis, especially in cases of resistance to commonly used pediculicides. However, as with other insecticides, IVM is subject to selection pressures that have led to the development of resistance in many arthropods. To gain insight into the mechanisms underlying IVM resistance in body lice, we both looked for mutations in the GluCl and searched the entire proteome for potential new loci involved in resistance. Firstly, the comparison of the cDNA GluCl between the Lab-IVR and Lab-IVS strains yielded no non-silent SNPs. This fact most likely excludes the involvement of GluCl in IVM resistance, at least under the conditions of the experiment. Furthermore, in most published studies that have implicated GluCl-target site insensitivity in IVM resistance, the level of 215 / 285 resistance of the organism was considerably high. This is the case of Plutella xylostella (~11 000-fold) [23], Tetranychus urticae (>2,000-fold) [24] and D. melanogaster (> 10-fold) [19]. Secondly, in our efforts to shed light on other possible mechanisms of resistance, we compared the overall protein expression pattern of the IVM-resistant and susceptible strains. In total, 22 proteins were differentially regulated, of which 13 were up-regulated and 9 were down-regulated in the resistant strain. Among the differentially expressed proteins found in the resistant strain, the most interesting observation was the activation of energy metabolism through the up-regulation of several key enzymes in the metabolic pathways (i.e. the isocitrate dehydrogenase [NAD] subunit, adenylate kinase, ATP synthase delta chain, tubulin alpha chain, tubulin beta chain, and ornithine aminotransferase), affecting the Krebs cycle, phosphorylation oxidative, purine, vitamins and amino acid metabolisms. Such nutrient availability may be necessary to overcome the elevated demands for energy and metabolism in the ‘toxic’ environment of the resistant lice, and consequently to maintain normal metabolism and energy balance at the cellular level. This is consistent with the fact that insecticide resistance is usually associated with higher demands for energy observed in other insect species [25,26]. A Cpx was the most significantly (P<0.0001) and highly downexpressed (10-fold) protein in the IVM-resistant lice. Indeed, this protein plays a key role in regulating synaptic exocytosis and neurotransmitter release. It was, therefore, selected to investigate its possible involvement in IVM-resistance. The Cpx cDNA from the body louse was identified and characterized. The deduced amino acid sequence presented very high similarity with the Cpx of other insects, suggesting that the basic mechanisms of its functions are similar to those described for D. melanogaster, the model organism, from which Cpx has been extensively studied. 216 / 285 The decrease of Cpx in protein expression was found to be associated with its mRNA downregulation, suggesting that the factors influencing Cpx expression occur at a pretranscriptional level. To gain insight into the mechanisms underlying this downexpression, we performed a DNA mutation analysis of Cpx transcripts and found that some mRNAs Cpx from the Lab-IVR strain had gained a premature stop codon. Therefore, the reduction of Cpx expression might be due, in part, to a secondary effect of a nonsense mutation inside the Cpx gene. Such a mechanism, known as nonsense-mediated mRNA decay, has been reported in insects [27,28] whereby mutations inside a gene cause premature termination codons and quickly degrade mRNA, inhibiting the accumulation of nonsense (inactive) proteins. RNAi techniques for the suppression of specific transcripts is proving to be a powerful tool in several insect species [29]. Body louse genome analysis has been shown to contain the genes necessary for RNAi [30]. Subsequent studies have reported that the injection of dsRNA can effectively suppress target genes in lice, and this ability has been widely used in gene function research [21,31]. Thus, we evaluated the resistance function of Cpx in susceptible lice via RNAi. Our findings showed that the injection of dsRNA-Cpx resulted in an effective suppression of Cpx expression and significantly decreased susceptibility to IVM compared with the control. This result provides convincing evidence that Cpx plays an important role in conferring IVM resistance in the body louse studied and represents the first evidence linking Cpx to insecticide resistance. Cpx is small neuron-specific cytosolic protein that interact with the assembled SNARE (soluble N ethylmaleimide-sensitive factor [NSF] attachment protein receptor) complex to modulate the vesicle fusion process and neurotransmitter release [22,32]. Despite intensive research, the precise functions of Cpx remain controversial and continue to present a conundrum [22,32]. For instance, genetic studies conducted in mammalians, worms and fruit flies have all shown that a Cpx has a dual function and can act either as inhibitory or 217 / 285 facilitatory for neurotransmitter release depending on the species, type of synapse (at CNS or at neuromuscular junction (NMJ)) and whether or not the vesicles are activated by Ca2+ (spontaneous or evoked release) [22]. Although it is not known how this downregulated protein is associated with IVM resistance, as IVM acts as a ligand for the inhibitory ligand- gated ion channel, activated by its natural agonist neurotransmitter glutamate [11,33], we hypothesized that the effect of Cpx on IVM resistance may be through regulating the glutamate release machinery at glutamatergic synapses. Indeed, studies conducted on worms have shown that the affinity of IVM is enhanced dramatically in the presence of glutamate, suggesting that the natural ligand, by binding to a distinct site, can allosterically enhance the activity of IVM and exert complementary, and possibly additive, effects on the conformational changes needed for the channels to open [33–35]. Moreover, it was thought that the extraordinary potency of the IVM killing parasites at much lower concentrations than those needed to activate recombinant channels expressed in Xenopus oocytes [34] is due to the interaction between endogenous glutamate and IVM [35]. Taken together, one could speculate that the downexpression of Cpx constitutes a primary mechanism by which lice protect their CNS, given that the GluCls in insects are mostly expressed in CNS [36] and the function of Cpx in that synapses is speculated to be mostly facilitatory for neurotransmitter release [22]. Although many issues remain to be investigated, the result of our study is exciting and provides the first insights into the mechanism underlaying IVM resistance in body lice at proteomic level, and links Cpx to insecticide resistance for the first time. 218 / 285 Experimental procedures 1. Ethics statement Adult New Zealand white rabbits were obtained from Charles River laboratories, were handled according to the rules of N° 2013-118, February 7, 2013, France and the experimental protocols (references APAFIS # 01077.02 & 2015050417122619), were approved by the Ethics Committee “C2EA-14” of Aix-Marseilles University, France and the French Ministry of National Education, Higher Education and Research. 2. Body louse populations Body louse strain and rearing. The Orlando (Culpepper) reference strain of body lice, P. h. humanus, was used in this study. The lice were maintained at 29°C with 70%–90% humidity and fed three times a week on the shaved abdomen of anesthetized rabbits. This strain has been reared in the insectary of our laboratory for more than fifteen years without exposure to any insecticides and is used as a reference susceptible strain referred to as Lab- IVS. Resistant population selection. The IVM resistant strain was selected from a susceptible strain with IVM, referred as Lab-IVR. The selection was performed by exposing each generation of adult lice to IVM for 30 minutes, by feeding them on a rabbit specifically dedicated to this purpose, that had received a subcutaneous injection of IVM (IVOMEC®, Merial) three hours previously. The experiments started by exposing the lice to an approximate median lethal dose (LD50) of 100 µg/kg of the rabbit’s body weight. This dose was determined through a preliminary assay conducted to generate dose response data (Table S1). After three generations, the selected strain was then exposed to the constant concentration of 150 µg/kg. This dose killed 90 to 100% of susceptible Lab-IVS. Selection was carried out over seven months and mortality was maintained at around 20–30%. Bioassays for IVM susceptibility was undertaken at the end of the selection. The LD50, the 219 / 285 median lethal times (LT50) and 95% confidence intervals were calculated by probit analysis using the SPSS software (IBM software, Armonk, NY). 3. Comparison of proteomic profiles between IVM-resistant and susceptible lice Protein preparation and digestion. Total protein was extracted from both IVM- resistant and susceptible lice (10 starved adult lice per sample, with four replicates of each). Samples were suspended in 200 μL of lysis buffer (8 M urea, 2 M thiourea, 100 mM NaCl, 25 Mm Tris, pH 8.2, complete protease inhibitor) and crushed with two 3-mm tungsten beads in TissueLyser at 25 Hz for four minutes (Qiagen, Courtaboeuf, France). After homogenization, the insoluble fractions were removed by centrifugation and soluble proteins were dialyzed twice using Slide-A-Lyzer MINI Dialysis Devices (Pierce Biotechnology, Rockford, USA) and dialysis buffer (50 mM ammonium bicarbonate, pH 7.4, 1 M Urea) following the manufacture’s protocol. Dialyzed fractions were collected, and protein concentration was determined by Bradford Protein Assay using Coomassie (Biorad, Marnes- la-Coquette, France). The dialyzed fractions were used as a template for global proteomic analysis. Briefly, 50 µg of total soluble proteins were reduced with 10 mM dithiothreitol for one hour at 30°C, alkylated with 20 mM iodoacetamide for one hour in the dark, and then digested by adding 2 µg of sequencing-grade trypsin solution (Promega, Charbonnières, France) for 20 hours at 37°C. The digested samples were then desalted using Pierce Detergent Removal Spin Columns (Thermo Fisher Scientific, Illkirch, France) following the manufacturer’s protocol. Label-free quantitative nano-LC-MS/MS proteomics analysis. The protein digests were analyzed using a NanoAcquity UPLC System with two-dimensional liquid chromatography (2D-LC) Technology (Waters, Saint-Quentin-en-Yvelines, France) connected to a Synapt G2Si Q-TOF ion mobility hybrid mass spectrometer (TWIM-MS; Waters, Saint-Quentin-en-Yvelines, France). The first chromatographic dimension consisted 220 / 285 of a 300-μm by 50-mm C18 column (Nano Ease 5 μm XBridge BEH130, Waters). Peptides were eluted onto a second dimension using a gradient of seven steps at 1.5 μl/min, with 20 mM ammonium formate pH 10, and 12, 15, 18, 20, 25, 35, and 65% acetonitrile. A trapping column (nanoAcquity UPLC 10K-2D-V/M Trap 5-μm Symmetry C18 column; 180 μm × 20 mm, Waters) was used to collect the first-dimension peptides for concentration and desalting, after dilution at 20 μl/min in 99.9% water–0.1% formic acid and 0.1% acetonitrile–0.1% formic acid. The second dimension consisted of a 75-μm by 250-mm C18 column (nanoAcquity UPLC 1.8-μm HSS T3; Waters). Peptides eluted from the first-dimension steps were separated using a 1-h gradient (275 nl/min; 5 to 40% acetonitrile–0.1% formic acid). Data-independent MS/MS analysis was performed with the ion mobility feature (HDMSe method). The ion source parameters were capillary voltage 3 kV, sampling cone voltage 40 V, ion source temperature 90°C, cone gas flow 50 L/h. Transfer collision low energy was set to 5 V and trap collision low energy was set to 4 V. The high energy ramp was applied from 4 V to 5 V for the trap collision and from 19 V to 45 V for the transfer collision enabling fragmentation of the ions after the ion mobility cell and before the time-of-flight (TOF) MS. On-column sample load was 285 ng per fraction (2 µl injected). Data processing and analysis. The acquired files were imported into Progenesis QI software Version 2.0 (Nonlinear Dynamics, Newcastle, UK) for label-free quantification analysis. The data were aligned automatically and normalized. Processing parameters were 150 counts for the low energy threshold, 30 counts for the elevated energy threshold. Databases used to identify peptides combined data from Phthiraptera (TrembL, 03/17/2016, 14,329 sequences) and the Oryctolagus (TrembL, 03/18/2016, 23,018 sequences). Search tolerance parameters were: peptide and fragment tolerance, 15 ppm, FDR < 1%; minimum ion matching requirements were three fragments per peptide, seven fragments per protein and two peptides per protein. The enzyme specificity was trypsin allowing one missed cleavage, 221 / 285 carbamidomethyl of cysteine (fixed), oxidation of methionine (variable), carbamyl of lysine and N-terminal (variable). Protein normalization was performed according to the relative quantitation using non-conflicting peptides. ANOVA tests were performed to determine the significance of changes between samples. A fold-change of >2 and a p-value <0.05 in at least two replicates were used as the thresholds to define differently expressed proteins. Gene ontology analysis for the differentially expressed proteins was carried out using Blast2GO (https://www.blast2go.com/blast2go-pro/download-b2g). The metabolic pathway analysis was conducted according to the Kyoto Encyclopedia of Genes and Genomes (KEGG) Pathway Database (http://www.genome.jp/kegg). 4. Total RNA extraction and reverse transcriptase-quantitative real-time PCR analysis (qRT-PCR) Isolation of RNA and cDNA synthesis. Total RNA from a pool of six lice from both the IVM-resistant and susceptible strains was extracted using the RNeasy Mini kit (Qiagen) according to the manufacturer’s instructions. The quantity and quality of the RNA were assessed using a NanoDrop ND-1000 (Thermo Fisher Scientific). First-strand cDNA was synthesized using MMLV-RT kit (Invitrogen) with oligo (dT) as primer, according to the manufacturer’s protocol. Quantification of mRNA expression by quantitative real time PCR (qPCR). Primers for qPCRs were designed from all the selected genes using the free web Primer3 software, version 4.0 (http://frodo.wi.mit.edu/primer3/) and their sequences were listed in Table S2. qPCRs were performed using a CFX96™ Real-Time system (Bio-Rad Laboratories, Foster City, CA, USA) with LightCycler® FastStart DNA Master SYBR Green I (Roche applied Science) in accordance with the manufacturer’s instructions. Three biological replicates, with three technical replications for each, were evaluated for each sample. We chose the 222 / 285 housekeeping gene elongation factor 1-α (EF1α) for internal normalization [21]. The Fold- changes (FC) of target genes relative to EF1α were calculated according to the 2−ΔΔCt method [37]. 5. Cloning and sequence analysis of the Cpx and GluCl transcripts Rapid amplification of cDNA ends (RACE) with SMARTer RACE cDNA Amplification kits (Clontech, PaloAlto, CA, USA) was used to obtain the full-length cDNA of Cpx following the manufacturer’s protocol, using universal primers supplied in the kits and gene-specific primers (GSPs) designed based on the partial cDNA sequence annotated from the body lice genome sequencing project (GenBank accession XM002426374). Subsequently, the full-length cDNA of Cpx was generated using a specific primer pair designed based on the 5’and 3’end sequences of the putative Cpx mRNA. Full-length cDNA was subjected to bioinformatic analysis using an ORF (open reading frame) finder tool (http://www.ncbi.nlm.nih.gov/gorf/gorf.html). Subsequently, the complete ORF of Cpx was amplified using the same cDNA synthesized for qRT-PCR from both resistant and susceptible strains to perform DNA polymorphism analysis. Amplification of the ORF cDNA GluCl was also conducted, as described for the Cpx gene, using a set of primers designed based on the cDNA gene sequence available in the NCBI database (GenBank accession XM002429761). All primers used are listed in Table S2. PCRs amplifications were performed using a Peltier PTC-200 thermal cycler (MJ Research Inc., Watertown, MA, USA) with the Hotstar Taq-polymerase (Qiagen). The purified PCR products were ligated into a pGEMT-easy vector (Promega) and transformed into JM109 Competent Cells. The plasmid inserts were PCR amplified using a vector-specific primer (M13 forward and reverse primers) and subjected to sequencing using the Big Dye Terminator Cycle Sequencing Kit (Perkin Elmer Applied Biosystems, Foster City, CA) with an ABI automated sequencer (Applied Biosystems). The electropherograms were assembled 223 / 285 using ChromasPro (ChromasPro 1.7, Technelysium Pty Ltd., Tewantin, Australia). Alignment of the nucleotide and amino-acid sequences was conducted using the ClustalW2 computer program (http://www.ebi.ac.uk/Tools/clustalw2/index.html) and phylogenetic trees were constructed with MEGA7.1. 6. Functional validation of the role of Cpx gene in IVM-resistance by RNAi dsRNA synthesis and gene knockdown. A Cpx transcription template from the cDNA of wild type lice was generated by PCR amplification using gene-specific primers with the T7 promoter element attached at their 5’ ends (Table S2). The purified PCR product was then used as a template in the dsRNA synthesis reaction using the MEGAscript RNAi Kit (Invitrogen) according to the manufacturer’s instructions. The dsRNA quality was evaluated by gel electrophoresis and quantified with a NanoDrop ND-1000 (Thermo Fisher Scientific). The dsRNA of the Escherichia coli plasmid amplified from pQE-30 vector (Qiagen) was prepared as described above and used as control. RNAi experiments were carried out by injecting ~120 ng of dsRNA-Cpx to adult lice in the ventral side between the second and third posterior abdominal segments, as described previously [21], with the FemtoJet 4i injector (Eppendorf, Germany), using self-pulled glass capillary needles (World Precision Instruments, Germany) under an Axio Zoom V16 microscope (Carl Zeiss S.A.S., France). The injected dose was determined empirically through preliminary experiments by determining concentrations and volumes of dsRNA resulting in maximum levels of target gene silencing that caused no mortality at various time post-injection. Lice were also injected with the dsRNA of pQE-30 plasmid as control. Total RNA was extracted from injected lice and processed to cDNA synthesis and qPCRs analysis to evaluate the degree of target gene silencing as described above. The relative expression level of Cpx of each ds-Cpx sample relative to the controls was calculated according to the 2−ΔΔCt method [37] after normalization with the EF1α gene. The Student’s t-test was used to assess the statistical significance using 224 / 285 GraphPad Prism version 7.00 for Windows (GraphPad Software, La Jolla California USA, www.graphpad.com). Mortality bioassays following dsRNA-Complexin injection. Two groups of lice from either the Cpx dsRNA-injected (165 lice) or pQE30 dsRNA-injected (150 lice), at 48 hours post-injection, were exposed to IVM by feeding them on a rabbit that had received a dose of 150 µg/kg of IVM. Mortality was assessed over six hours after the IVM-treatment to calculate the value of LT50 and RR, as described above. The Chi-2 test was used to assess the statistical significance. 225 / 285 Acknowledgements We thank Pascal Weber for helping to pull glass capillary needles, and to Anne-Marie Gottrau and Jean-Michel Berenger for help maintaining the body lice rearing system. This study was supported by the Fondation Méditerranée Infection and the French National Research Agency under the “Investissements d’avenir” program, reference ANR-10-IAHU- 03. 226 / 285 References 1. Bonilla DL, Durden LA, Eremeeva ME, Dasch GA. The biology and taxonomy of head and body lice--implications for louse-borne disease prevention. PLoS Pathog. 2013;9: e1003724. doi:10.1371/journal.ppat.1003724 2. Chosidow O. Scabies and pediculosis. The Lancet. 2000;355: 819–826. doi:10.1016/S0140-6736(99)09458-1 3. Veracx A, Raoult D. Biology and genetics of human head and body lice. Trends Parasitol. 2012;28: 563–571. doi:10.1016/j.pt.2012.09.003 4. Raoult D, Roux V. The body louse as a vector of reemerging human diseases. Clin Infect Dis Off Publ Infect Dis Soc Am. 1999;29: 888–911. doi:10.1086/520454 5. Brouqui P. Arthropod-Borne Diseases Associated with Political and Social Disorder. Annu Rev Entomol. 2011;56: 357–374. doi:10.1146/annurev-ento-120709-144739 6. Amanzougaghene N, Fenollar F, Sangaré AK, Sissoko MS, Doumbo OK, Raoult D, et al. Detection of bacterial pathogens including potential new species in human head lice from Mali. PLOS ONE. 2017;12: e0184621. doi:10.1371/journal.pone.0184621 7. Raoult D, Abat C. Developing new insecticides to prevent chaos: the real future threat. Lancet Infect Dis. 2017;17: 804–805. doi:10.1016/S1473-3099(17)30395-X 8. Durand R, Bouvresse S, Berdjane Z, Izri A, Chosidow O, Clark JM. Insecticide resistance in head lice: clinical, parasitological and genetic aspects. Clin Microbiol Infect Off Publ Eur Soc Clin Microbiol Infect Dis. 2012;18: 338–344. doi:10.1111/j.1469-0691.2012.03806.x 9. Chosidow O, Giraudeau B, Cottrell J, Izri A, Hofmann R, Mann SG, et al. Oral ivermectin versus malathion lotion for difficult-to-treat head lice. N Engl J Med. 2010;362: 896–905. doi:10.1056/NEJMoa0905471 10. Chaccour C, Hammann F, Rabinovich NR. Ivermectin to reduce malaria transmission I. Pharmacokinetic and pharmacodynamic considerations regarding efficacy and safety. Malar J. 2017;16: 161. doi:10.1186/s12936-017-1801-4 11. Laing R, Gillan V, Devaney E. Ivermectin - Old Drug, New Tricks? Trends Parasitol. 2017;33: 463–472. doi:10.1016/j.pt.2017.02.004 12. Ōmura S, Crump A. Ivermectin and malaria control. Malar J. 2017;16. doi:10.1186/s12936-017-1825-9 13. Leulmi H, Diatta G, Sokhna C, Rolain J-M, Raoult D. Assessment of oral ivermectin versus shampoo in the treatment of pediculosis (head lice infestation) in rural areas of Sine-Saloum, Senegal. Int J Antimicrob Agents. 2016;48: 627–632. doi:10.1016/j.ijantimicag.2016.07.014 14. Mumcuoglu KY, Miller J, Rosen LJ, Galun R. Systemic activity of ivermectin on the human body louse (Anoplura: Pediculidae). J Med Entomol. 1990;27: 72–75. 227 / 285 15. Pariser DM, Meinking TL, Bell M, Ryan WG. Topical 0.5% ivermectin lotion for treatment of head lice. N Engl J Med. 2012;367: 1687–1693. doi:10.1056/NEJMoa1200107 16. Chen L-P, Wang P, Sun Y-J, Wu Y-J. Direct interaction of avermectin with epidermal growth factor receptor mediates the penetration resistance in Drosophila larvae. Open Biol. 2016;6. doi:10.1098/rsob.150231 17. Gao X, Yang J, Xu B, Xie W, Wang S, Zhang Y, et al. Identification and Characterization of the Gene CYP340W1 from Plutella xylostella and Its Possible Involvement in Resistance to Abamectin. Int J Mol Sci. 2016;17. doi:10.3390/ijms17030274 18. Diatta G, Abat C, Sokhna C, Tissot-Dupont H, Rolain J-M, Raoult D. Head lice probably resistant to ivermectin recovered from two rural girls in Dielmo, a village in Sine-Saloum, Senegal. Int J Antimicrob Agents. 2016;47: 501–502. doi:10.1016/j.ijantimicag.2016.03.013 19. Kane NS, Hirschberg B, Qian S, Hunt D, Thomas B, Brochu R, et al. Drug-resistant Drosophila indicate glutamate-gated chloride channels are targets for the antiparasitics nodulisporic acid and ivermectin. Proc Natl Acad Sci U S A. 2000;97: 13949–13954. doi:10.1073/pnas.240464697 20. Luo L, Sun Y-J, Wu Y-J. Abamectin resistance in Drosophila is related to increased expression of P-glycoprotein via the dEGFR and dAkt pathways. Insect Biochem Mol Biol. 2013;43: 627–634. doi:10.1016/j.ibmb.2013.04.006 21. Yoon KS, Strycharz JP, Baek JH, Sun W, Kim JH, Kang JS, et al. Brief exposures of human body lice to sub-lethal amounts of ivermectin over transcribes detoxification genes involved in tolerance. Insect Mol Biol. 2011;20: 687–699. doi:10.1111/j.1365- 2583.2011.01097.x 22. Trimbuch T, Rosenmund C. Should I stop or should I go? The role of complexin in neurotransmitter release. Nat Rev Neurosci. 2016;17: 118–125. doi:10.1038/nrn.2015.16 23. Wang X, Wang R, Yang Y, Wu S, O’Reilly AO, Wu Y. A point mutation in the glutamate-gated chloride channel of Plutella xylostella is associated with resistance to abamectin. Insect Mol Biol. 2016;25: 116–125. doi:10.1111/imb.12204 24. Dermauw W, Ilias A, Riga M, Tsagkarakou A, Grbić M, Tirry L, et al. The cys-loop ligand-gated ion channel gene family of Tetranychus urticae: Implications for acaricide toxicology and a novel mutation associated with abamectin resistance. Insect Biochem Mol Biol. 2012;42: 455–465. doi:10.1016/j.ibmb.2012.03.002 25. Silva AX, Jander G, Samaniego H, Ramsey JS, Figueroa CC. Insecticide Resistance Mechanisms in the Green Peach Aphid Myzus persicae (Hemiptera: Aphididae) I: A Transcriptomic Survey. PLOS ONE. 2012;7: e36366. doi:10.1371/journal.pone.0036366 228 / 285 26. Ahmad M, Denholm I, Bromilow RH. Delayed cuticular penetration and enhanced metabolism of deltamethrin in pyrethroid-resistant strains of Helicoverpa armigera from China and Pakistan. Pest Manag Sci. 2006;62: 805–810. doi:10.1002/ps.1225 27. Long AA, Mahapatra CT, Woodruff EA, Rohrbough J, Leung H-T, Shino S, et al. The nonsense-mediated decay pathway maintains synapse architecture and synaptic vesicle cycle efficacy. J Cell Sci. 2010;123: 3303–3315. doi:10.1242/jcs.069468 28. Casadio A, Longman D, Hug N, Delavaine L, Vallejos Baier R, Alonso CR, et al. Identification and characterization of novel factors that act in the nonsense-mediated mRNA decay pathway in nematodes, flies and mammals. EMBO Rep. 2015;16: 71–78. doi:10.15252/embr.201439183 29. Abd El Halim HM, Alshukri BMH, Ahmad MS, Nakasu EYT, Awwad MH, Salama EM, et al. RNAi-mediated knockdown of the voltage gated sodium ion channel TcNav causes mortality in Tribolium castaneum. Sci Rep. 2016;6: 29301. doi:10.1038/srep29301 30. Pittendrigh BR, Olds BP, Yoon KS, Lee SH, Sun W, Clark JM. The genomics of human lice: From the genome to the potential for future control strategies. Pestic Biochem Physiol. 2013;106: 172–176. doi:10.1016/j.pestbp.2013.03.010 31. Kwon DH, Kim JH, Kim YH, Yoon KS, Clark JM, Lee SH. Identification and characterization of an esterase involved in malathion resistance in the head louse Pediculus humanus capitis. Pestic Biochem Physiol. 2014;112: 13–18. doi:10.1016/j.pestbp.2014.05.006 32. Huntwork S, Littleton JT. A complexin fusion clamp regulates spontaneous neurotransmitter release and synaptic growth. Nat Neurosci. 2007;10: 1235–1237. doi:10.1038/nn1980 33. Forrester SG, Beech RN, Prichard RK. Agonist enhacement of macrocyclic lactone activity at a glutamate-gated chloride channel subunit from Haemonchus contortus. Biochem Pharmacol. 2004;67: 1019–1024. doi:10.1016/j.bcp.2003.08.047 34. Pemberton DJ, Franks CJ, Walker RJ, Holden-Dye L. Characterization of Glutamate- Gated Chloride Channels in the Pharynx of Wild-Type and Mutant Caenorhabditis elegansDelineates the Role of the Subunit GluCl-α2 in the Function of the Native Receptor. Mol Pharmacol. 2001;59: 1037–1043. doi:10.1124/mol.59.5.1037 35. Wolstenholme AJ, Rogers AT. Glutamate-gated chloride channels and the mode of action of the avermectin/milbemycin anthelmintics. Parasitology. 2005;131: S85–S95. doi:10.1017/S0031182005008218 36. Kita T, Ozoe F, Azuma M, Ozoe Y. Differential distribution of glutamate- and GABA- gated chloride channels in the housefly Musca domestica. J Insect Physiol. 2013;59: 887–893. doi:10.1016/j.jinsphys.2013.06.005 37. Schmittgen TD, Livak KJ. Analyzing real-time PCR data by the comparative CT method. Nat Protoc. 2008;3: nprot.2008.73. doi:10.1038/nprot.2008.73 229 / 285 Figure 1. Comparison of proteomic and qRT-PCR results. The x-axis shows the 15 selected genes, while the y-axis gives the fold change observed for the Lab-IVR vs the Lab- IVS strains. EF1α was used to normalize the mRNA levels. Values are means ±SEMs (n=3). The selected genes are: Cpx: Complexin, Tryp: Trypsin, CLTC: Clathrin heavy chain, RPS3A: 40S ribosomal protein S3a, TUBα1: Tubulinα1, OAT: Ornithine aminotransferase, HSP: Heat shock protein, Idh: Isocitrate dehydrogenase subunit, Adk: Adenylate kinase, ATPase: ATP synthase, Fib-H: Heavy-chain filboin, MO-porin: Mitochondrial outer membrane porin channel, Lmpt: Limpet, Tubß2: Tubulinß2 and E0W0W7. 230 / 285 Figure 2. Comparison of Cpx proteins. (A) Phylogenetic tree showing the phylogenetic relationships of Cpx genes from Insecta and non-Insecta species. The tree was generated by ClustalW alignment of the amino acid sequences of Cpx genes using the neighbor-joining (NJ) method. (B) Amino acid sequence alignment of P. humanus (Ph) with members of the Cpx family. Identical residues are marked green and highly conserved residues are marked red. The blue box indicates the position of the predicted SNARE-binding domain. Dm: Drosophila melanogaster; Ce: Caenorhabditis elegans, Cc: Cyphomyrmex costatus, Ag: Anoplophora glabripennis. 231 / 285 Figure 3. The effect of dsRNA on expression of Cpx in the IVM-susceptible strain and the effect of Cpx knockdown on IVM resistance. (A) Cpx mRNA levels were quantified by qRT-PCR in 72 hours of injection of dsRNA Cpx or pQE30 (control). The change in mRNA levels in the ds-RNA Cpx were calculated relative to controls. Values are means +SEMs (n=3). Asterisks (*) indicate that Cpx dsRNA significantly suppresses the levels of Cpx transcript (t-test, *P<0.05 and **P<0.01). (B) Bioassays for Lab-IVS lice exposed to IVM (150 µg/Kg) started 48 hours post-injection of dsRNA Cpx or pQE30 (control). Asterisks (*) indicate that knocking down Cpx expression decreased IVM-susceptibility in the Lab-IVS strain compared to the control (Chi-2 test, *P<0.05 and **P<0.01). 232 / 285 Table 1. Resistance of the selected strain (Lab-IVR) to IVM based on a comparison of median lethal (LT50) at a dose of 150 µg/Kg. Lice N LT50 (hours) (95% CL) Slope ± SE χ2 RR Lab-IVS 500 28.83 (24.47-32.78) 4 ± 0.85 10 - Lab-IVR (selected) 300 157.01 (144.91-172.37) 0.86± 0.061 3.4 5.4 CL: Confidence limited; SE: Standard error; RR: Resistance ratio calculated by dividing the LT 50 of the Lab-IVR strain by LT50 of Lab-IVS strain 233 / 285 Table 2. Differentially expressed proteins identified by proteomic analysis of the IVM- resistant strain compared to the susceptible strain. Acc. ID Description Peptide Unique Anova Fold count Peptide (p) change Up-regulated E0W486 Adenylate kinase 16 15 0.000 3.08 E0VX56 ATP synthase delta chain, putative 11 11 0.005 2.23 E0W1N3 Heavy-chain filboin, putative 6 6 0.006 2.51 E0VW06 Mitochondrial outer membrane porin channel 28 25 0.009 2.12 E0VGH4 Limpet, putative 7 7 0.009 4.14 E0VSM7 Tubulin alpha-1 chain 75 19 0.015 3.05 E0VGF0 Putative uncharacterized protein 10 9 0.018 2.16 E0VQ79 Tubulin beta-2 chain, putative 17 8 0.024 2.31 E0W0W7 Putative uncharacterized protein 4 3 0.029 3.85 E0VYE0 Ornithine aminotransferase, putative 6 6 0.034 2.10 E0VHY3 Heat shock protein, putative 6 6 0.043 2.03 Isocitrate dehydrogenase [NAD] subunit, 7 7 0.048 E0VSN4 2.61 mitochondrial E0W2K0 Ejaculatory bulb-specific protein 3, putative 51 16 0.049 3.37 Down-regulated E0VJX5 Complexin, putative 3 3 0.000 -10.24 Sodium/potassium-transporting ATPase subunit 3 2 0.004 E0W229 -2.97 beta-2, putative E0VPU2 40S ribosomal protein S3a 3 2 0.004 -3.93 E0VWY3 D-beta-hydroxybutyrate dehydrogenase, putative 5 4 0.009 -2.68 E0VD43 Clathrin heavy chain 19 16 0.02 -3.24 E0VFA6 Trypsin 4 3 0.03 -3.78 E0VQK9 Guanine nucleotide-binding protein G(O) subunit 6 2 0.035 -2.85 alpha, putative E0VRZ6 Zinc finger protein CDGSH domain-containing 3 3 0.035 -4.69 protein, putative E0VKP4 Actin, muscle 164 61 0.05 -2.31 234 / 285 Table 3. Functional annotation of differentially expressed proteins. Acc. No. Gene name GO analysis GO analysis GO analysis InterPro (IP) KEGG_PATHWAY (Molecular function) (Biological process) (Cellular component) Adenylate kinase activity, ADP biosynthetic process, mitochondrial Adenylate kinase Purine metabolism, Thiamine Adenylate E0W486 ATP binding, AMP metabolic process, ATP intermembrane space, metabolism, Biosynthesis of kinase, putative metabolic process, cytosol, antibiotics E0VX56 ATP synthase proton-transporting ATPase ATP synthesis coupled proton proton-transporting ATP ATPase, F1 Oxidative phosphorylation, delta chain, activity, rotational mechanism transport synthase complex, catalytic complex, Purine metabolism, Thiamine putative core delta/epsilon metabolism subunit E0W1N3 Heavy-chain filboin, putative E0VW06 Mitochondrial voltage-gated anion channel sperm individualization, mitochondrial outer Porin 235 /285 outer membrane activity transmembrane transport, membrane, microtubule porin channel regulation of cilium assembly, associated complex, lipid regulation of anion particle, nebenkern transmembrane transport, mitochondrial transport, photoreceptor cell maintenance E0VGH4 limpet, putative zinc ion binding Zinc finger, LIM-type, E0VSM7 Tubulin alpha GTP binding, GTPase microtubule-based process cytoplasm, microtubule Purine metabolism, Thiamine chain activity, structural constituent metabolism of cytoskeleton E0VGF0 Uncharacterized methyltransferase activity, methylation Farnesoic acid protein transferase activity O-methyl transferase E0VQ79 Tubulin beta GTP binding, GTPase microtubule-based process cytoplasm, microtubule Purine metabolism, Thiamine chain activity, structural constituent metabolism of cytoskeleton E0W0W7 Uncharacterized carboxypeptidase activity proteolysis membrane, integral carboxypeptidase protein component of membrane inhibitor E0VYE0 Ornithine pyridoxal phosphate binding, arginine catabolic process to cytoplasm Arginine and proline aminotransferase identical protein binding, glutamate, arginine catabolic metabolism, Metabolic ornithine-oxo-acid process to proline via pathways, Biosynthesis of transaminase activity ornithine antibiotics E0VHY3 Heat shock Stress response protein, putative E0VSN4 Isocitrate magnesium ion binding, tricarboxylic acid cycle mitochondrion, integral Isocitrate and Citrate cycle (TCA cycle), dehydrogenase isocitrate dehydrogenase component of membrane isopropylmalate Metabolic pathways, [NAD] subunit, (NAD+) activity, NAD dehydrogenases Biosynthesis of antibiotics, mitochondrial binding family Carbon metabolism, 2- Oxocarboxylic acid metabolism, Biosynthesis of amino acids E0W2K0 Ejaculatory Insect bulb-specific pheromone- 236 /285 protein 3 binding protein precursor A10/OS-D, E0VJX5 Complexin, syntaxin binding neurotransmitter transport Synaphin, putative E0W229 Sodium potassium ion transport, sodium: potassium- potassium- sodium ion transport exchanging ATPase transporting complex ATPase subunit beta-2 E0VPU2 40S ribosomal structural constituent of translation cytosolic small ribosomal Ribosomal Ribosome protein S3a ribosome subunit protein S3Ae E0VWY3 D-beta- retinol dehydrogenase activity oxidation-reduction process integral component of Glucose/ribitol hydroxybutyrate membrane dehydrogenase, dehydrogenase NAD(P)-binding domain, E0VD43 Clathrin heavy structural molecule activity vesicle-mediated transport, Clathrin coat of trans-Golgi Lysosome, Endocytosis chain intracellular protein transport network vesicle, Clathrin coat of coated pit E0VFA6 Trypsin serine-type endopeptidase proteolysis Neuroactive ligand-receptor activity interaction E0VQK9 Guanine GTP binding, G-protein adenylate cyclase-modulating heterotrimeric G-protein nucleotide- coupled receptor binding, G-protein coupled receptor complex binding protein metal ion binding, GTPase signaling pathway G(O) subunit activity, G-protein alpha, beta/gamma-subunit complex binding, signal transducer activity, metal ion binding E0VRZ6 Zinc finger 2 iron, 2 sulfur cluster intracellular membrane- protein CDGSH binding bounded organelle, integral domain- component of membrane containing protein E0VKP4 Actin, muscle ATP binding, isopentenyl- isoprenoid biosynthetic Biosynthesis of antibiotics, diphosphate delta-isomerase process Terpenoid backbone activity, hydrolase activity biosynthesis 237 /285 Table 4. Comparison of median lethal time (LT50) between the lice with its Cpx knockdown (Cpx dsRNA-injected) and control lice (pQE30 dsRNA-injected). Lice N LT50 (h) (95% CL) Slope ± SE χ2 RR Cpx dsRNA-injected 165 92.54 (81.90-105.07) 0.78 ± 0.07 6.33 3.22 pQE30 dsRNA-injected 150 27.69 (24.31-30.82) 4± 0.11 1.94 0.96 RR: Resistance ratio calculated by dividing the LT50 of either the Cpx dsRNA or pQE30 dsRNA by LT 50 of Lab-IVS 238 / 285 Supporting Information Table S1. Susceptibility of the Lab-IVS strain to IVM. Probit regression data for the relationship between dose of ivermectin and mortality at 72 hours. Lice LD50 (µg/kg) (95% CL) LD90 (µg/kg) (95% CL) Slope ± SE χ2 S-Lab 98.36 (83.56-114.67) 152.59 (132.76-187.76) 3 ± 0.01 5.8 CL: Confidence limited; SE: Standard error; LD: Lethal dose 239 / 285 Table S2. Primer sequences used in this study. Amplification Gene name Forward Sequence (3’-5’) Reverse Sequence (3’-5’) of primes qPCR analysis Complexin ACTGTAGCACGCATTTGCTTTC GCCGCAAGAAGAACCAAAT Trypsin GCCGGATCATCCAAAGCTAA TGAGTCCAACAACGGGTTTG Clathrin heavy GCCGTCAAAGCCGATAGAAC CTTTCGGCAAATTCGTAGGC chain 40S ribosomal GGCGGTCGGTAAGAATAAGG CCCTCGGATGCAATTTTTGT Tubulinα1 GAACTGTCGCCAAATTACTTCC CGCTTGCTGTTTGTTATACAGG Ornithine CTCGGCAAATACGATTCTAGCT CATCCCCGTATCGTAAACGTA T Heat shock CAAACCGTCGTACCAAGTGTTA CTCACACATCGGAAACAACAT protein C Isocitrate AGGTGCTGG TGTTGTCGCG ATTGAGCATTTTGGCGGCGC dehydrogenase Adenylate CTCGGTAACATCGGAGGA GGGTGAAGACGCTGAAAAA kinase ATP synthase GCTTCTGCAACTTCTACAGCAA CCTGGAGTTGTAACCGTTTTTC Heavy-chain ACCAAATCCAACAGCTGTACCT CACCTTCCACAGGTGGATTT filboin Mitochondrial GTGAGATTTTGACCTTCCGAAC TCACATCAAGACAAGGGTTCA porin G Limpet CACAATATGGCTTCTCATCTCG GACTCGTTGCGTCAAGTGTAA C Tubulinß2 TCCCTCTGCTTCTTTTCTAACG GGTACAATGGATTCCGTACGT T E0W0W7 ACCGAGACCAGAACAAGATGT CCGGAAAACAGTTGCAGTAAT EF1α CGTTTCCGTAAAAGAATTGCG GGCTATGTGAGCCGTATGAC dsRNA Complexin X.ATGCCGCAAGAAGAACCAA X.TCACTGTAGCACGCATTT G pQE30 X.GTTCATCCATAGTTGCCTGAC X.AGATAACACTGCGGCCAACT T TAC RACE Complexin- CTTCCTCAGCTTCCGCTTCCGCT GSP-5’ GC Complexin- GCAGCGGAAGCGGAAGCTGAG GSP-3’ Full-length Complexin ATGGCGGCTTTCGTTGCA TCACTGTAGCACGCATTTGC amplification GluCl ATGGAATACAGCGTTCAGCT TTAATTATCTTCCGTATCTTCT C X: T7 promoter (TAATACGACTCACTATAGGG); GSP: Gene specific primer 240 / 285 Table S3. Cpx accession numbers from distinct species used for construction of phylogenetic tree. Species list Cpx Accession no. Vertebrates Homo sapiens AHW56455 Narke japonica O42105 Rattus norvegicus NM_022864 Xenopus laevis NP_001087909 Bos taurus AAI22584 Nematodes Trichuris trichiura CDW57795 Strongyloides ratti CEF68549 Caenorhabditis elegans NP_490868 Arthropods Ixodes scapularis EEC07951 Daphnia pulex EFX87986 Aedes aegypti XP021712522 Drosophila melanogaster NM001170033 Cyphomyrmex costatus XP018396052 Anoplophora_glabripennis XM018709052 Stomoxys calcitrans XP013102718 241 / 285 Figure S1. Gene ontology (GO) analysis of the differentially expressed proteins. The proteins are grouped into three GO terms: cellular component, biological process and molecular function. 242 / 285 Figure S2. Gene ontology assignment of downregulated and upregulated proteins related to molecular function (MF), biological processes (BP) and cellular component (CC). 243 / 285 Figure S3. Amino acid sequences alignment of body lice Cpx transcripts. The alignment shows a frameshift starting at amino acid 100 and a premature stop codon at amino acid 111 in the mutated transcript (from the resistant lice) compared to normal Cpx transcript. 244 / 285 Conclusions et perspectives Le travail présenté dans ce manuscrit s’inscrit dans la continuité des nombreux travaux réalisés en sein de l’IHU-méditerranée infection (ex-URMITE) sur le pou humain où nous avons voulu approfondir les connaissances et répondre à certaines questions concernant les poux humains restées en débat depuis longtemps. Ainsi, dans la première partie de ce travail nous avons mis en évidence pour la première fois la présence de clade B au Moyen-Orient, datant de plus de 2.000 ans, supportant une origine asiatique pour ce clade, suivi par son introduction dans le Nouveau Monde avec les premiers hommes ayant atteint le continent américain il y a des milliers d’années. En outre, nous avons mis en évidence l’existence d’un nouveau clade mitochondrial (clade F) en Amazonie, indiquant que la diversité génétique chez le pou est plus élevée que l'on ne le pensait auparavant. Dans les études futures, il serait très intéressant de séquencer le maximum de poux appartenant à différents clades et provenant de différents endroits à travers le monde pour effectuer une analyse globale des génomes. L'intégration des profils phylogénomiques et génomiques des populations permettra d'avoir des informations plus complètes sur l'évolution des poux mais aussi sur notre histoire évolutive. Dans la deuxième partie de ce travail, nous avons développé une PCR en temps réel pour l’identification de routine rapide des clades de poux. Cet outil s’est révélé très utile sur le terrain notamment pour l’analyse d’une large collection de poux provenant de différents pays, ce qui nous a permis d’avoir une idée plus claire sur la distribution des clades. De plus, nous avons mis en évidence la présence d’ADN de plusieurs bactéries dans les poux de tête dont certaines, telles que C. burnetii, R. aeschlimannii, B. theileri, Ehrlichia et Anaplasma, ont été détectées pour la premier fois chez les poux. L’ensemble des résultats obtenus renforcent l’hypothèse d’un rôle vectoriel probable du pou de tête et soulève la nécessité d’approfondir les recherches 245 / 285 à ce propos. Des études futures devraient viser à mieux comprendre les différents facteurs qui peuvent influencer la différence dans la capacité vectorielle observée entre les poux de corps et les poux de tête, notamment les interactions existantes entre les communautés microbiennes présentes au sein des poux (en particulier le rôle des bactéries endosymbiotiques). Des technologies de séquençage à haut débit pourrait être envisagées afin de réaliser une exploration exhaustive des symbiotes et pathogènes des poux. Enfin, le travail présenté dans cette thèse apporte des données nouvelles sur la résistance des poux à l’ivermectine, en mettant en évidence la présence de trois mutations non-synonymes. Ces mutations pouvant induire un changement conformationnel qui induirait une diminution de l’affinité de l’ivermectine pour le canal GluCl. Afin de confirmer ceci, une validation fonctionnelle en utilisant des lignées de poux transgéniques exprimant des variants de GluCl porteurs de ces trois mutations, suivie par une exposition à l’ivermectine est nécessaire. En parallèle, grâce à notre nouveau système de détection moléculaire par PCR-RFLP il sera possible de suivre l’évolution de ces mutations sur le terrain. L’exposition répétée d’une population de poux de corps de laboratoire (la souche Orlando) à des doses subléthales répétées d’ivermectine à plusieurs générations a permis, pour la première fois à notre connaissance, de sélectionner une population de poux résistante à l’ivermectine. Une analyse protéomique globale comparative entre les poux résistants et les poux sensibles a révélé qu’une répression significative de la complexine pourrait être à l’origine de la résistance à l’ivermectine chez ces poux. Le rôle de la complexine a été par la suite clairement confirmé par une validation fonctionnelle par les ARN interférents. Cette découverte représente la première évidence liant la complexine à la résistance aux insecticides. Afin de compléter cette étude de laboratoire, d’autres expérimentations pourraient être réalisées. Dans un premier temps, il serait intéressant de poursuivre la sélection de la population de poux résistante à l’ivermectine afin de suivre l’évolution de niveau de l’expression de la 246 / 285 complexine en fonction de l’évolution de la résistance des poux. Une caractérisation fonctionnelle plus approfondie, en utilisant des approches plus avancées, est nécessaire pour élucider le mécanisme exact par lequel la complexine induit la résistance, une question essentielle à laquelle nous n’avons pas pu répondre pour le moment. Il serait aussi intéressant d’explorer le rôle des autres protéines différemment transcrites que nous avons identifié au cours de notre étude en plus de la complexine, et qui semblent également susceptibles d’induire la résistance. Bien qu'aucune mutation non-synonyme au niveau de GluCl n’a été détectée chez les poux-résistants sélectionnés au laboratoire, présentant un niveau faible de résistance, nous pensons que la poursuite de la sélection permettra de mieux caractériser ce gène ainsi que les mécanismes par lesquels il induit la résistance à l’ivermectine. Par la suite, il serait intéressant de savoir si les mécanismes identifiés chez ces poux résistants de laboratoire existent sur des poux sauvages. Enfin, il n’en demeure pas moins intéressant d’appliquer ces approches d’exploration globale, telles que la protéomique et transcriptomique, afin d’explorer et de mieux comprendre les résistances chez les poux vis-à-vis d’autres insecticides, dont les mécanismes moléculaires sont très peu connus (comme ceux vis-à-vis du malathion et de la perméthrine) voire complétement inconnus (comme ceux vis-à-vis des organophosphorés, du carbaryl et du lindane). Ceci est primordial afin de préserver les rares insecticides encore disponibles, en attendant la mise sur le marché de nouvelles molécules plus spécifiques ou de nouveaux moyens de lutte plus efficaces contre les poux ainsi que les maladies qu’ils transmettent. 247 / 285 Références bibliographiques 1. Reed DL, Light JE, Allen JM, 2008;197: 535–543. doi:10.1086/526520 Kirchman JJ. Pair of lice lost or parasites regained: the evolutionary 8. Reed DL, Smith VS, Hammond SL, history of anthropoid primate lice. Rogers AR, Clayton DH. Genetic BMC Biol. 2007;5: 7. Analysis of Lice Supports Direct doi:10.1186/1741-7007-5-7 Contact between Modern and Archaic Humans. PLOS Biol. 2004;2: e340. 2. Drali R, Abi-Rached L, Boutellis A, doi:10.1371/journal.pbio.0020340 Djossou F, Barker SC, Raoult D. Host switching of human lice to new world 9. Boutellis A, Drali R, Rivera MA, monkeys in South America. Infect Mumcuoglu KY, Raoult D. Evidence of Genet Evol J Mol Epidemiol Evol sympatry of clade a and clade B head Genet Infect Dis. 2016;39: 225–231. lice in a pre-Columbian Chilean doi:10.1016/j.meegid.2016.02.008 mummy from Camarones. PloS One. 2013;8: e76818. 3. Veracx A, Raoult D. Biology and doi:10.1371/journal.pone.0076818 genetics of human head and body lice. Trends Parasitol. 2012;28: 563–571. 10. Ascunce MS, Fane J, Kassu G, Toloza doi:10.1016/j.pt.2012.09.003 AC, Picollo MI, González-Oliver A, et al. Mitochondrial diversity in human 4. Light JE, Allen JM, Long LM, Carter head louse populations across the TE, Barrow L, Suren G, et al. Americas. Am J Phys Anthropol. Geographic distributions and origins of 2013;152: 118–129. human head lice (Pediculus humanus doi:10.1002/ajpa.22336 capitis) based on mitochondrial data. J Parasitol. 2008;94: 1275–1281. 11. Boutellis A, Bitam I, Fekir K, Mana N, doi:10.1645/GE-1618.1 Raoult D. Evidence that clade A and clade B head lice live in sympatry and 5. Drali R, Shako J-C, Davoust B, Diatta recombine in Algeria. Med Vet G, Raoult D. A New Clade of African Entomol. 2015;29: 94–98. Body and Head Lice Infected by doi:10.1111/mve.12058 Bartonella quintana and Yersinia pestis- Democratic Republic of the Congo. Am 12. Sunantaraporn S, Sanprasert V, J Trop Med Hyg. 2015;93: 990–993. Pengsakul T, Phumee A, Boonserm R, doi:10.4269/ajtmh.14-0686 Tawatsin A, et al. Molecular survey of the head louse Pediculus humanus 6. Ashfaq M, Prosser S, Nasir S, Masood capitis in Thailand and its potential role M, Ratnasingham S, Hebert PDN. High for transmitting Acinetobacter spp. diversity and rapid diversification in the Parasit Vectors. 2015;8: 127. head louse, Pediculus humanus doi:10.1186/s13071-015-0742-4 (Pediculidae: Phthiraptera). Sci Rep. 2015;5. doi:10.1038/srep14188 13. Boutellis A, Veracx A, Angelakis E, Diatta G, Mediannikov O, Trape J-F, et 7. Raoult D, Reed DL, Dittmar K, al. Bartonella quintana in head lice from Kirchman JJ, Rolain J-M, Guillen S, et Sénégal. Vector Borne Zoonotic Dis al. Molecular identification of lice from Larchmt N. 2012;12: 564–567. pre-Columbian mummies. J Infect Dis. doi:10.1089/vbz.2011.0845 248 / 285 14. Sangaré AK, Boutellis A, Drali R, 22. Kim JH, Previte DJ, Yoon KS, Murenzi Socolovschi C, Barker SC, Diatta G, et E, Koehler JE, Pittendrigh BR, et al. al. Detection of Bartonella quintana in Comparison of the proliferation and African body and head lice. Am J Trop excretion of Bartonella quintana Med Hyg. 2014;91: 294–301. between body and head lice following doi:10.4269/ajtmh.13-0707 oral challenge. Insect Mol Biol. 2017;26: 266–276. 15. Kittler R, Kayser M, Stoneking M. doi:10.1111/imb.12292 Molecular evolution of Pediculus humanus and the origin of clothing. 23. Angelakis E, Rolain J-M, Raoult D, Curr Biol CB. 2003;13: 1414–1417. Brouqui P. Bartonella quintana in head louse nits. FEMS Immunol Med 16. Toups MA, Kitchen A, Light JE, Reed Microbiol. 2011;62: 244–246. DL. Origin of Clothing Lice Indicates doi:10.1111/j.1574- Early Clothing Use by Anatomically 695X.2011.00804.x Modern Humans in Africa. Mol Biol Evol. 2011;28: 29–32. 24. Piarroux R, Abedi AA, Shako J-C, doi:10.1093/molbev/msq234 Kebela B, Karhemere S, Diatta G, et al. Plague epidemics and lice, Democratic 17. Raoult D, Roux V. The body louse as a Republic of the Congo. Emerg Infect vector of reemerging human diseases. Dis. 2013;19: 505–506. Clin Infect Dis Off Publ Infect Dis Soc Am. 1999;29: 888–911. 25. Raoult D, Abat C. Developing new doi:10.1086/520454 insecticides to prevent chaos: the real future threat. Lancet Infect Dis. 18. Raoult D. A Personal View of How 2017;17: 804–805. doi:10.1016/S1473- Paleomicrobiology Aids Our 3099(17)30395-X Understanding of the Role of Lice in Plague Pandemics. Microbiol Spectr. 26. Bonilla DL, Durden LA, Eremeeva ME, 2016;4. Dasch GA. The biology and taxonomy doi:10.1128/microbiolspec.PoH-0001- of head and body lice--implications for 2014 louse-borne disease prevention. PLoS Pathog. 2013;9: e1003724. 19. Angelakis E, Diatta G, Abdissa A, doi:10.1371/journal.ppat.1003724 Trape J-F, Mediannikov O, Richet H, et al. Altitude-dependent Bartonella 27. Durand R, Bouvresse S, Berdjane Z, quintana genotype C in head lice, Izri A, Chosidow O, Clark JM. Ethiopia. Emerg Infect Dis. 2011;17: Insecticide resistance in head lice: 2357–2359. clinical, parasitological and genetic doi:10.3201/eid1712.110453 aspects. Clin Microbiol Infect Off Publ Eur Soc Clin Microbiol Infect Dis. 20. Boutellis A, Mediannikov O, Bilcha 2012;18: 338–344. doi:10.1111/j.1469- KD, Ali J, Campelo D, Barker SC, et al. 0691.2012.03806.x Borrelia recurrentis in head lice, Ethiopia. Emerg Infect Dis. 2013;19: 28. Chosidow O, Giraudeau B, Cottrell J, 796–798. doi:10.3201/eid1905.121480 Izri A, Hofmann R, Mann SG, et al. Oral ivermectin versus malathion lotion 21. Murray ES, Torrey SB. Virulence of for difficult-to-treat head lice. N Engl J Rickettsia prowazeki for head lice. Ann Med. 2010;362: 896–905. N Y Acad Sci. 1975;266: 25–34. doi:10.1056/NEJMoa0905471 249 / 285 29. Diatta G, Abat C, Sokhna C, Tissot- Dupont H, Rolain J-M, Raoult D. Head lice probably resistant to ivermectin recovered from two rural girls in Dielmo, a village in Sine-Saloum, Senegal. Int J Antimicrob Agents. 2016;47: 501–502. doi:10.1016/j.ijantimicag.2016.03.013 30. Kirkness EF, Haas BJ, Sun W, Braig HR, Perotti MA, Clark JM, et al. Genome sequences of the human body louse and its primary endosymbiont provide insights into the permanent parasitic lifestyle. Proc Natl Acad Sci U S A. 2010;107: 12168–12173. doi:10.1073/pnas.1003379107 31. Laing R, Gillan V, Devaney E. Ivermectin - Old Drug, New Tricks? Trends Parasitol. 2017;33: 463–472. doi:10.1016/j.pt.2017.02.004 32. Chosidow O, Giraudeau B, Cottrell J, Izri A, Hofmann R, Mann SG, et al. Oral ivermectin versus malathion lotion for difficult-to-treat head lice. N Engl J Med. 2010;362: 896–905. doi:10.1056/NEJMoa0905471 33. Diatta G, Abat C, Sokhna C, Tissot- Dupont H, Rolain J-M, Raoult D. Head lice probably resistant to ivermectin recovered from two rural girls in Dielmo, a village in Sine-Saloum, Senegal. Int J Antimicrob Agents. 2016;47: 501–502. doi:10.1016/j.ijantimicag.2016.03.013 250 / 285 Annexes 251 / 285 Article 11 : Multiple Pathogens Including Potential New Species in Tick Vectors in Côte d'Ivoire Publié dans PLoS Negl Trop Dis. 2016; 15:e0004367 252 / 285 RESEARCH ARTICLE Multiple Pathogens Including Potential New Species in Tick Vectors in Côte d’Ivoire Cyrille Bilé Ehounoud1,2,3, Kouassi Patrick Yao3, Mustapha Dahmani1, Yaba Louise Achi4, Nadia Amanzougaghene1,Adèle Kacou N’Douba5, Jean David N’Guessan3, Didier Raoult1,2, Florence Fenollar1,2, Oleg Mediannikov1,2* 1 Aix-Marseille Université, URMITE, UM63, CNRS 7278, IRD 198, Inserm U1095, Faculté de médecine, Marseille cedex 05, France, 2 Campus International UCAD-IRD, Dakar, Senegal, 3 Felix Houphouet Boigny Université, UFR Biosciences, Côte D’Ivoire, 4 Ecole de spécialisation en Elevage de Bingerville, Côte D’Ivoire, 5 Felix Houphouet Boigny Université, UFR Sciences médicales, Côte D’Ivoire Abstract OPEN ACCESS Background Citation: Ehounoud CB, Yao KP, Dahmani M, Achi Our study aimed to assess the presence of different pathogens in ticks collected in two YL, Amanzougaghene N, Kacou N’Douba A, et al. regions in Côte d’Ivoire. (2016) Multiple Pathogens Including Potential New Species in Tick Vectors in Côte d’Ivoire. PLoS Negl Trop Dis 10(1): e0004367. doi:10.1371/journal. Methodology/Principal Findings pntd.0004367 Real-time PCR and standard PCR assays coupled to sequencing were used. Three hun- Editor: Joseph M. Vinetz, University of California, dred and seventy eight (378) ticks (170 Amblyomma variegatum, 161 Rhipicepalus micro- San Diego School of Medicine, UNITED STATES plus,3Rhipicephalus senegalensis,27Hyalomma truncatum,16Hyalomma marginatum Received: November 9, 2015 rufipes, and 1 Hyalomma impressum) were identified and analyzed. We identified as patho- Accepted: December 15, 2015 genic bacteria, Rickettsia africae in Am. variegatum (90%), Rh. microplus (10%) and Hya- Published: January 15, 2016 lomma spp. (9%), Rickettsia aeschlimannii in Hyalomma spp. (23%), Rickettsia massiliae in Rh. senegalensis (33%) as well as Coxiella burnetii in 0.2%, Borrelia sp. in 0.2%, Anaplasma Copyright: © 2016 Ehounoud et al. This is an open access article distributed under the terms of the centrale in 0.2%, Anaplasma marginale in 0.5%, and Ehrlichia ruminantium in 0.5% of all Creative Commons Attribution License, which permits ticks. Potential new species of Borrelia, Anaplasma, and Wolbachia were detected. Candi- unrestricted use, distribution, and reproduction in any datus Borrelia africana and Candidatus Borrelia ivorensis (detected in three ticks) are phylo- medium, provided the original author and source are genetically distant from both the relapsing fever group and Lyme disease group borreliae; credited. both were detected in Am. variegatum. Four new genotypes of bacteria from the Anaplas- Data Availability Statement: All relevant data are mataceae family were identified, namely Candidatus Anaplasma ivorensis (detected in within the paper. three ticks), Candidatus Ehrlichia urmitei (in nine ticks), Candidatus Ehrlichia rustica (in four Funding: This study was funded by the IHU ticks), and Candidatus Wolbachia ivorensis (in one tick). Méditerranée Infection (http://en.mediterranee- infection.com/), UEMOA (PASRES Project) (http:// www.csrs.ch/pasres/) and the association APRI Conclusions/Significance (Marseille, France). The funders had no role in study design, data collection and analysis, decision to For the first time, we demonstrate the presence of different pathogens such as R. aeschli- publish, or preparation of the manuscript. mannii, C. burnetii, Borrelia sp., A. centrale, A. marginale, and E. ruminantium in ticks in Competing Interests: The authors have declared Côte d’Ivoire as well as potential new species of unknown pathogenicity. that no competing interests exist. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 1 / 18 253 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Author Summary The management of febrile illnesses represents a veritable challenge in sub Saharan-Africa. Until recently most of them were considered as malaria. However, it was showed that a large part of non-malarial febrile diseases in African rural regions (for instance, in Senegal) may be caused by tick-borne infections. Unfortunately, no data exist about the prevalence and incidence of tick-borne diseases in Côte d'Ivoire and their role in public health. We aimed to search for different pathogenic bacteria in ticks in order to understand if there is the background for tick-borne diseases. We detected pathogenic bacteria responsible for many infectious diseases like Rickettsia (spotted fevers), Borrelia (relapsing fevers), Ana- plasma, Ehrlichia (ehrlichiosis and anaplasmosis) and Coxiella burnetii (Q fever). These finding suggested that, as in others sub-Saharan African countries, tick-borne disease may be considered as a health care problem in Cote d'Ivoire. Introduction Ticks are important vectors of many pathogens and are considered as the second biggest vec- tors of human and animal diseases after mosquitoes [1,2]. Many tick-borne bacterial emerging diseases such as spotted fevers, borrelioses, anaplasmoses, ehrlichioses, and Q fever have been described worldwide [3,4,5]. It was recently shown that in many tropical countries tick- and acari-borne infections play important role in human pathology. In Senegal, for instance, arthropod-borne borreliosis and rickettsiosis were identified in 16.3% of acute fevers recorded by rural dispensaries [6]. Acari-borne tsutsugamushi fever is one of the major causes of acute febrile morbidity in South-Eastern Asia [7]. Investigations of the vectors of tick-borne diseases are one of the main keys to controlling related morbidity [8]. Rickettsioses, caused by bacteria belonging to the spotted fever group (SFG) of the genus Rickettsia, are considered among the oldest known vector-borne zoonotic diseases [9]. The most common rickettsia in Africa is Rickettsia africae, the etiological agent of African tick- borne fever [10]. This disease has been reported with high seroprevalence in sub-Saharan Afri- can countries including Cameroon (11.9% - 51.8%) and Senegal (21.4% - 51%) [11,12]. R. afri- cae has been detected by PCR in ticks in Mali, Niger, Burundi, and Sudan [13]. Amblyomma hebraeum and Amblyomma variegatum ticks are the main reservoirs and vectors of R. africae in Southeastern Africa and sub-Saharan Africa, respectively [9,14]. It was also reported in other species of Amblyomma such as Amblyomma lepidum in Djibouti [15] and Amblyomma compressum in the Democratic Republic of Congo and Liberia [16,17]. In Western Africa, R. africae has been detected in several Rhipicephalus ticks including Rhipicephalus annulatus in Guinea, Senegal, and Nigeria [12,16,18], Rhipicephalus evertsi evertsi in Senegal and Nigeria [12,18], Rhipicephalus decoloratus in Nigeria [19], Rhipicephalus geigyi in Liberia [16], and Hyalomma spp. ticks including Hyalomma impeltatum in Nigeria [18] and Hyalomma margin- atum rufipes in Guinea [16] but not in Côte d’Ivoire, where a strain of R. africae has been iso- lated from Am. variegatum [20]. Rickettsia aeschlimannii is an agent of spotted fever which was first identified in a patient returning from Morocco [21]. In this country, it was first isolated from Hyalomma margina- tum marginatum ticks [22]. R. aeschlimannii was also reported by PCR in other Hyalomma ticks including H. marginatum rufipes and Hyalomma truncatum ticks collected from camels and cows in Egypt, Algeria, Sudan, and Tunisia [23]. In Western Africa, R. aeschlimannii was also detected in 15% to 95% of H. marginatum rufipes from Mali, Niger, Senegal and Nigeria [12,13,24] and in 6% to 7% of H. truncatum from Senegal [12] but not in Côte d’Ivoire. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 2 / 18 254 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Rickettsia massiliae is another SFG rickettsia. Since its description in 2005, R. massiliae infec- tions in humans have been confirmed in Europe and South America [25,26,27]. It is associated with Rhipicephalus ticks. R. massiliae was found by PCR in Rhipicephalus spp. ticks including Rhipicephalus spp. from Côte d’Ivoire [28], Rhipicephalus guilhoni from Senegal [12], Rhipice- phalus senegalensis from Guinea [16], and Rhipicephalus eversti from Nigeria [18]. Different borrelioses are caused by bacteria from the Borrelia genus. They are traditionally classified into the Lyme disease group and the relapsing fever group. The former is ecologically associated with hard ticks and is mostly found in the temperate northern hemisphere [29]. Relapsing fever group borreliae are mostly associated with soft ticks and found in subtropical regions worldwide [30,31]. In endemic regions, borrelioses may play an important role, for example in Slovakia [32]. Relapsing fever is one of the most common diseases in several Afri- can regions including Senegal [33,34] and east African countries [35]. It is caused by different Borrelia species such as Borrelia hispanica, Borrelia duttonii, and Borrelia crocidurae. B. hispa- nica was recently detected in 11.6% to 20% of Ornithodoros ticks from northern Africa [31,36]. B. crocidurae is responsible for tick-borne relapsing fever in West Africa. Its distribution in the south is thought to be limited by the 750 mm isohyets [37]. Neither this borrelia nor any other from the relapsing group has been reported in Côte d’Ivoire. A controversial study, based on molecular data, reported 30 cases of borreliosis in Togo but its epidemiology was not identified [38] and studies in neighboring countries did not confirm the presence of borreliosis in west tropical sub-Saharan Africa. In Ethiopia, Borrelia sp. was recently identified by PCR in 7.3% of Amblyomma cohaerens [39]. Phylogenetically, this Borrelia sp. was placed in an intermediate position between Lyme disease and relapsing fever groups. All bacteria from the Anaplasmataceae family are intracellular mammal parasites, arthro- pods nematodes, and trematodes [40]. Anaplasma centrale and Anaplasma marginale are two etiological agents of bovine anaplasmosis in ruminants [41]. These species are distributed in tropical and subtropical regions of Africa and naturally infect cattle [42]. They were previously found by molecular biology in ticks in neighboring Mali [43]. These bacteria are often found in Dermacentor, Rhipicephalus, and Amblyomma ticks throughout the world [44]. Ehrlichia rumi- nantium is responsible for cowdriosis in ruminants with the Amblyomma genus ticks as a vec- tor [45]. Cowdriosis induces mortality in ruminants in sub-Saharan Africa and in islands in the Caribbean where it causes serious losses to animal production [40]. E. ruminantium was previously identified in Am. variegatum in Burkina Faso but not in Côte d’Ivoire. No cases of human ehrlichiosis or anaplasmosis have been reported in Africa, but recently human patho- gens such as Anaplasma phagocytophilum have been reported in Senegal and Algeria [46,47]. Bacteria from the Wolbachia genus of the Anaplasmatacae family are associated with arthro- pods and filarial nematodes. They are responsible for reproductive alterations in arthropods which are indirectly (via nematodes) associated with human pathogenesis [40]. Finally, Q fever is a zoonotic disease caused by Coxiella burnetii. This bacterium may cause severe infections such as chronic endocarditis and abortion [48,49]. It infects humans usually by a direct contact with domestic animals such as cattle, sheep, goats, and dogs [50]. It was pre- viously reported in Amblyomma, Rhipicephalus, and Dermacentor ticks [43]. In Senegal, C. burnetii was detected in 0.8% to 14.2% of ticks including Am. variegatum, Rhipicephalus spp., Hyalomma spp., and Ornithodoros sonrai [51] and may play a role in Q fever epidemiology. In Côte d’Ivoire, the seroprevalence was estimated at 3.4% [52]. Although these diseases have emerged in many African countries, they remain neglected. In Côte d’Ivoire, little information is available about these diseases and their epidemiology. To date, the existence and/or prevalence of tick-borne associated pathogens remain poorly under- stood. Our study provides the first data screening for multiple tick-borne associated pathogens in Côte d’Ivoire. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 3 / 18 255 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Materials and Methods Ethics statement To perform this study, an approval of Cote d'Ivoire Ethics committee was received under the number N°86/MSLS/CNERN-dkn. Period, study area and tick collection The tick collection was conducted over a period ranging from October 30 to November 8, 2014. Ticks were manually collected from cattle in two regions of Côte d'Ivoire: Savannah and Bandama Valley (Fig 1, Table 1). In total, 378 ticks (304 adults and 74 nymphs) were collected Fig 1. Map of Côte d’Ivoire showing the regions, cities and villages where the ticks were collected for our study. doi:10.1371/journal.pntd.0004367.g001 PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 4 / 18 256 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Table 1. Geographic coordinates of tick collection sites. City1 or village2 Geographic coordinates Species Number (male/female/nymphs) Total number Savannah region Kong1 08°15N 05°07W Rh. microplus 3/10/0 13 Ferké1 09°55N 05°20W Am. variegatum 7/3/4 14 Rh. microplus 1/4/0 5 H. impressum 1/0/0 1 H. truncatum 0/1/0 1 Kafolo-bac2 09°43N 04°39W Am. variegatum 1/0/5 6 Rh. microplus 3/8/1 12 H. truncatum 1/0/0 1 Torokinkenin2 08°84N 04°47W Am. variegatum 2/2/1 5 Rh. microplus 4/10/0 14 Téhini2 09°60N 03°67W Am. variegatum 2/1/0 3 Rh. microplus 1/4/1 6 H. marginatum rufipes 0/1/0 1 Doropo2 09°77N 03°40W Am. variegatum 2/0/9 11 Rh. microplus 0/7/0 7 H. marginatum rufipes 2/2/0 4 H. truncatum 0/2/0 2 Naissain2 09°39N 04°48W Am. variegatum 0 /1/4 5 Rh. microplus 1/4/0 5 Sikolo2 09°43N 04°66W Am. variegatum 0/0/4 4 Rh. microplus 1/4/0 5 Fasselemon2 09°27N 04°52W Am. variegatum 4/1/0 5 Rh. microplus 1/4/0 5 Nafana2 09°18N 04°78W Am. variegatum 1/0/0 1 Rh. microplus 1/1/0 2 Tindara2 09°54N 04°74W Am. variegatum 4/2/11 17 Rh. microplus 0/12/0 12 H. truncatum 1/0/0 1 Laleraba2 10°13N 05°08W Am. variegatum 4/1/0 5 Rh. microplus 0/5/0 5 H. truncatum 7/3/0 10 H. marginatum rufipes 1/2/0 3 Hamdalaye2 09°98N 05°13W Am. variegatum 10/6/3 19 Rh. microplus 1/4/0 5 H. truncatum 7/2/0 9 H. marginatum rufipes 2/1/0 3 Monogon2 09°82N 04°92 W Am. variegatum 2/3/0 5 Rh. microplus 2/4/0 6 H. truncatum 1/2/0 3 H. marginatum rufipes 2/3/0 5 Bandama Valley region Dabakala1 08°36N 04°41W Am. variegatum 10/2/2 14 Rh. microplus 2/4/0 6 Katiola1 08°15N 05°07W Am. variegatum 3/0/4 7 Rh. microplus 1/4/0 5 Niakara1 06°60N 05°29W Am. variegatum 8/2/9 19 (Continued) PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 5 / 18 257 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Table 1. (Continued) City1 or village2 Geographic coordinates Species Number (male/female/nymphs) Total number Rh. microplus 3/2/0 5 Darala2 08°44N 04°35W Am. variegatum 2/1/0 3 Rh. microplus 1/4/0 5 Rh. senegalensis 1/1/0 2 Ouandarama2 08°68N 04°39W Am. variegatum 2/0/4 6 Rh. microplus 1/4/1 6 Rh. senegalensis 0/1/0 1 Darakokaha2 08°27N 05°16W Am. variegatum 1/0/0 1 Rh. microplus 1/4/0 5 N’golodougou2 09°15N 05°12W Am. variegatum 4/2/0 6 Rh. microplus 2/7/0 9 Kolokaha2 08°97N 05°21W Am. variegatum 2/2/0 4 Rh. microplus 1/4/0 5 Sépikaha2 08°91N 05°03W Am. variegatum 1/0/9 10 Rh. microplus 2/4/2 8 Petionara2 08°47N 05°03W Rh. microplus 1/4/0 5 Am. variegatum 72/29/69 170 Rh. microplus 34/122/5 161 Rh. senegalensis 1/2/0 3 H. truncatum 17/10/0 27 H. marginatum rufipes 7/9/0 16 H. impressum 1/0/0 1 Total of ticks for the two regions 132/172/74 378 doi:10.1371/journal.pntd.0004367.t001 from three cities and 12 villages in the Savannah region and three cities and seven villages in the Bandama Valley region (Table 1). Ticks were stored in 70% ethanol until morphological and molecular analyses in laboratory of URMITE, Marseille (France). The species and sex of the ticks were identified according to standard taxonomic keys for adult ticks [2]. DNA extraction and real-time PCR Total DNA from half of each tick was extracted using the EZ1 DNA tissue kit (Qiagen, Hilden, Germany) following the manufacturer’s instructions. DNA extracts were stored at +4°C until use. Bacterial DNA was initially detected using bacterial genus-specific or species-specific quantitative real-time PCRs (qPCRs) targeting: Rickettsia spp., R. africae, R. aeschlimannii, R. massiliae, Borrelia spp., Anaplasmataceae spp., A. phagocytophilum, Bartonella spp., C. burne- tii, and Spiroplasma spp. (Table 2). Samples with a high discordance in the cycle threshold number (Ct) for Rickettsia spp. and R. africae (in all cases, low Ct for Rickettsia spp. and high Ct for R. africae) were subjected to specific qPCRs for two other rickettsial species: R. aeschli- mannii and R. massiliae in order to identify possible co-infection. qPCRs were performed using a CFX 96 Real Time System (Bio-Rad, Marnes-la-Coquette, France) and the Eurogentec MasterMix Probe PCR kit (Eurogentec, Liège, Belgium). PCR tests were considered to be posi- tive when the Ct was lower than 35 Ct [22]. In addition, two different specific qPCRs targeting two different sequences had to be positive in order to confirm the presence of a bacterium in the ticks. Positive controls (bacterial DNA) and negative controls (master mix or water) were used to validate the PCR runs. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 6 / 18 258 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Standard PCR and sequencing Most of samples which were considered positive by qPCRs were subsequently subjected to standard PCR. All samples which were positive using Rickettsia genus-specific but negative with R. africae qPCR were subjected to standard PCR to amplify a portion of the ompA gene. We also chose two positive ticks for R. africae by species to confirm the presence of R. africae by standard PCR. The primers used (190.70, 190.180, and 190.701) amplified a 632-bp frag- ment of the Rickettsia ompA gene [60]. For the identification of Borrelia species, primers target- ing a portion of the flaB gene were used [33]. Anaplasmataceae spp. (Anaplasma spp., Ehrlichia spp., and Wolbachia spp.) were identified using Ana 212f and Ana 753r primers tar- geting a 500 bp portion of the 23S rRNA gene [47]. Table 2. Primers and probes used for real-time quantitative PCR in this study. Microorganisms Targeted sequence Primers f, r (5’-3’) and Probes p (6FAM–TAMRA) References Rickettsia spp. gltA (RKNDO3) f_GTGAATGAAAGATTACACTATTTAT [53] r_GTATCTTAGCAATCATTCTAATAGC p_CTATTATGCTTGCGGCTGTCGGTTC R. africae poT15-dam2 f_TGCAACACGAAGCACAAAAC [6] r_CCTCTTGCGAAACTCTACTT p_TGA CGTGTGGATTCGAGCACCGGA R. aeschlimannii Intergenic spacer (RaescSca1) f_AAAGAAATGGATTTCACGGCGAA [12] r_ACCAAGTAAACGTCTCGTAC p_TGGGGAAATATGCCGTATACGCAAGC R. massiliae Hypothetical protein f_CCAACCTTTTGTTGTTGCAC [54] r_TTGGATCAGTGTGACGGACT p_CACGTGCTGCTTATACCAGCAAACA Anaplasma spp. 23S rRNA (TtAna) f_TGACAGCGTACCTTTTGCAT [47] r_TGGAGGACCGAACCTGTTAC p_GGATTAGACCCGAAACCAAG Anaplasma phagocytophilum apaG f_TAAGCGCAGTTGGAAGATCA [55] r_CGGCACATCCACATAAAACA p_TGATGAACGGCTGGTATCAG Spiroplasma rpoB f_TGTTGGACCAAACGAAGTTG [55] r_CCAACAATTGGTGTTTGTGG p_GCTAACCGTGCTTTAATGGG Coxiella burnetii Insertion Sequence (IS1111) f_CAAGAAACGTATCGCTGTGGC [56] r_CACAGAGCCACCGTATGAATC p_CCGAGTTCGAAACAATGAGGGCTG (IS30A) f_CGCTGACCTACAGAAATATGTCC [57] r_GGGGTAAGTAAATAATACCTTCTGG p_CATGAAGCGATTTATCAATACGTGTATG Bartonella spp. Internal transcribed spacer16S (BartoITS3) f_GATGCCGGGGAAGGTTTTC [58] r_GCCTGGGAGGACTTGAACCT p_GCGCGCGCTTGATAAGCGTG Borrelia spp Internal transcribed spacer 16S RNA (Bor ITS4) f_GGCTTCGGGTCTACCACATCTA [59] r_CCGGGAGGGGAGTGAAATAG p_TGCAAAAGGCACGCCATCACC (Bor_16S) f_AGCCTTTAAAGCTTCGCTTGTAG [34] r_GCCTCCCGTAGGAGTCTGG p_CCGGCCTGAGAGGGTGAACGG doi:10.1371/journal.pntd.0004367.t002 PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 7 / 18 259 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Standard PCR was performed on a ThermalCycler (Applied Biosystem, Paris, France). The reactions were carried out using the Hotstar Taq-polymerase (Qiagen), in accordance with the manufacturer’s instructions. The amplicons were visualized using electrophoresis on a 1.5% agarose gel stained with ethidium bromide and examined using an ultraviolet transilluminator. The PCR products were purified using a PCR filter plate Millipore NucleoFast 96 PCR kit fol- lowing the manufacturer’s recommendations (Macherey–Nagel, Düren, Germany). The ampli- cons were sequenced using the BigDye Terminator Cycle Sequencing Kit (Applied Biosystems) with an ABI automated sequencer (Applied Biosystems).The sequences which were obtained were assembled using ChromasPro software (ChromasPro 1.7, Technelysium Pty Ltd.,Tewan- tin, Australia) and compared with those available in GenBank by NCBI BLAST (http://blast. ncbi.nlm.nih.gov/Blast.cgi). Phylogenetic analysis DNA sequences alignment was carried out using MEGA 6 (http://www.megasoftware.net/ mega.php). We selected the Bayesian method [61] using TOPALi 2.5 software (Biomathemat- ics and Statistics Scotland) to construct phylogenetic trees. Results Of the 378 ticks identified, 170 Am. variegatum, 161 Rh. microplus,3Rh. senegalensis,27H. truncatum,16H. marginatum rufipes, and one H. impressum were analyzed. No A. phagocyto- philum, Bartonella spp. and Spiroplasma spp. were detected in ticks. Rickettsia spp. was found in 187 of 378 ticks (49%); most of them, 174/378 (46%), were identified as R. africae with spe- cific qPCR (Table 3). R. africae was detected in 154/170 (90%) Am. variegatum, 16/161 (10%) Rh. microplus, 2/16 (12%) H. marginatum rufipes, 1/27 (4%) H. truncatum and 1/1 H. impres- sum (Table 3). To confirm the presence of R. africae, we performed standard PCR using two positive ticks per species. The BLAST search of the ompA gene sequences from ticks revealed 100% nucleotide identity with the ompA gene of R. africae detected in Am. variegatum col- lected in Antigua (GenBank EU622980). We amplified the ompA fragment in all ticks positive for Rickettsia spp. but negative for R. africae qPCR. The BLAST analyses showed that ompA sequences of R. aeschlimannii were detected in 7/16 (44%) H. marginatum rupifes and 3/27 (11%) H. truncatum. The sequences were 99% identical to those of R. aeschlimannii, previously detected in H. impeltatum collected in Egypt (GenBank HQ335157) and 100% identical to those detected in H. marginatum in Turkey (GenBank KF791251). R. massiliae was observed in 1/3 (33%) Rh. senegalensis with 100% similarity R. massiliae, previously detected in Rh. sene- galensis in Guinea (GenBank JN043508). Finally, these results were confirmed by a specific qPCR for R. aeschlimannii and R. massiliae (Table 2). We also performed these species-specific qPCR on three samples (two H. marginatum rufipes and one Rh. senegalensis) where we observed a high discordance (more than 5 Cts) between Rickettsia genus-specific qPCR (low Ct) and R. africae species-specific qPCR (higher Ct). We found that in all three cases, a co- infection by two rickettsia species: R. massiliae plus R. africae in Rh. senegalensis and R. aeschli- mannii plus R. africae in H. marginatum rufipes. C. burnetii was detected in one tick (Table 3). Screening of all ticks for Borrelia spp. using qPCR, detected 16/378 (4%) positive ticks. We succeeded in amplifying a fragment of flaB gene and 16S rRNA sequence only in 4/378 (1%) ticks. A BLAST search showed that these sequences probably belong to an undescribed species, because only 87% (288/329 bp), 87% (287/328 bp), 97% (319/328 bp), and 87% (288/329 bp) similarities were observed with, respectively, the flaB gene of Borrelia duttonii (GenBank AB105132), Borrelia sp. IA-1 (GenBank EU492387), Borre- lia sp. BrFlab (GenBank EF141022), and Borrelia sp. IA-1 (GenBank EU492387). The PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 8 / 18 260 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Table 3. Prevalence of positive ticks by PCR. Bacterium% Am. variegatum Rh. microplus Rh. senegalensis H. truncatum H. marginatum H. impressum Total (positives/tested) Rickettsia spp. 90% (154/170) 10% (16/161) 33% (1/3) 15% (4/27) 69% (11/16) 100% (1/1) 49% (187/378) R. africae 90% (154/170) 10% (16/161) - 4% (1/27) 12% (2/16) 100% (1/1) 46% (174/378) R. aeschlimannii - - - 11% (3/27) 44% (7/16) - 2% (10/378) R. massiliae - - 33% (1/3) - - - 0.2% (1/378) C. burnetii 0.6% (1/170) - - - - - 0.2% (1/378) Borrelia spp. 6% (11/170) 2% (3/161) - 4% (1/27) 6% (1/16) - 5% (16/378) Borrelia sp. genotype - 0.6% (1/161) - - - - 0.2% (1/378) TCI301 Candidatus Borrelia 1% (2/170) - - - - - 0.5% (2 /378) ivorensis Candidatus Borrelia 0.6% (1/170) - - - - - 0.2% (1/378) africana Anaplasma spp. 12% (21 /170) 24% (39/161) - 11% (3/27) - - 16% (63/378) Anaplasma centrale 0.6% (1/170) - - - - - 0.2% (1/378) Anaplasma marginale - 1% (2/161) - - - - 0.5% (2/378) Candidatus Anaplasma 2% (3/170) - - - - - 0.8% (3/378) ivorensis Ehrlichia sp. 3% (6/170) 3% (5/161) - 7% (2/27) - - 3% (13/378) Candidatus Ehrlichia 0.6% (1/170) 1% (2 /161) - 4% (1/ 27) - - 1% (4/378) rustica Candidatus Ehrlichia 3% (5/170) 2% (3/161) - 4% (1/27) - - 2% (9/378) urmitei Ehrlichia ruminantium 1% (2/170) - - - - - 0.5% (2/378) Candidatus Wolbachia - 0.6% (1 /161) - - - - 0.2% (1/378) ivorensis R. africae +R. - - - - 12% (2/16) - 0.5% (2/378) aeschlimannii R. africae +R. massiliae - - 33% (1/3) - - - 0.2% (1/378) R. africae +C. burnetii 0.6% (1/170) - - - - - 0.2% (1/378) R. africae +Borrelia sp. - 0.6% (1/161) - - - - 0.2% (1/378) R. africae + Candidatus 2% (3/170) - - - - - 0.8% (3/378) Borrelia Africana + Candidatus Borrelia ivorensis R. africae +Anaplasma 0.6% (1/170) - - - - - 0.2% (1/378) centrale R. africae +Anaplasma ------0.2% (1/378) marginale R. africae +Candidatus 1% (2/170) 0.6% (1/161) - - - - 0.8% (3/378) Anaplasma ivorensis R. africae + Candidatus 1% (2/170) - - - - - 0.5% (2/378) Ehrlichia urmitei = 0%; the name ‘Candidatus’ is employed here for the new species because they are not isolated doi:10.1371/journal.pntd.0004367.t003 phylogenetic position of this Borrelia is shown in Fig 2. Because these potentially new species had not previously been isolated, we propose the provisional names Candidatus Borrelia afri- cana for the genotype TCI22 and Candidatus Borrelia ivorensis for the genotypes TCI140 and TCI351. In a phylogenetic tree based on a 344 bp fragment of the Borreliae flaB gene, the sequences of Candidatus Borrelia africana and Candidatus Borrelia ivorensis are situated in the PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 9 / 18 261 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Borrelia genus near Borrelia sp. from Ethiopian Amblyomma cohaerens (GenBank JX089967) and are closer to the relapsing fever group than to that of Lyme disease. As previously shown, Ethiopian Borrelia group together with these new genotypes to form a separate and Fig 2. flaB gene-based phylogenetic analysis of the strains identified in this study. Phylogenetic tree highlighting the position of Borrelia sp. identified in the present study relative to borrelia type strains and uncultured borreliae. The flaB sequences were aligned using CLUSTALW, and phylogenetic inferences were obtained from a Bayesian phylogenetic analysis with the HKY+Γ; JC+ Γ and HKY+ Γ substitution models for the first, second and third codons respectively. The GenBank accession numbers are indicated at the end. Sequences obtained in the present study are in bold. The numbers at the nodes are the bootstrap values obtained by repeating the analysis 100 times to generate a majority consensus tree. There were a total of 300 positions in the final dataset. The scale bar indicates a 10% nucleotide sequence divergence. doi:10.1371/journal.pntd.0004367.g002 PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 10 / 18 262 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Fig 3. 23S rRNA based phylogenetic analysis of strains identified in this study. Phylogenetic tree highlighting the position of Anaplasma sp, Ehrlichia sp and Wolbachia sp identified in the present study relative to Anaplama, Ehrlichia and Wolbachia type and uncultured strains. The 23S rRNA sequences were aligned using MEGA 6 and phylogenetic inferences were obtained from a Bayesian phylogenetic analysis with the HKY standard model. doi:10.1371/journal.pntd.0004367.g003 well-supported (bootraps 100) branch on the phylogenetic tree situated between Lyme disease and relapsing fever clusters, albeit closer to the latter. We also identified Borrelia sp. (genotype TCI301) in Rh. microplus which was almost identical to Borrelia sp. previously identified in the same ticks in Brazil (GenBank EF141022). Sixty-three ticks were positive using qPCR targeting the 23S rRNA of Anaplasmataceae. Only 39 DNA samples were positive using qPCR were successfully amplified in standard PCR. A possible explanation may consist of the lower sensitivity of standard PCR compared to qPCR. After sequencing, we obtained good quality sequences for only 22 samples (22/378; 6%). We suggest that the poor sequence quality may be explained by co-infection by two or more species belonging to the Anaplasmataceae family. We have identified one case of A. centrale in Am. variegatum (100% identity with the A. centrale strain Israel, NR_076686), and two cases of E. ruminantium in Am. variegatum (100% identity with the E. ruminantium strain Welgevon- den, NR_077000). We have identified A. marginale in two Rh. microplus (100% of homology with A. marginale strain Florida, NR_0765879). Finally, for all remaining sequences, Blast anal- ysis shows a homology score of under 92% which means that these sequences are likely to PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 11 / 18 263 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Table 4. New sequences amplified in this study and deposited in GenBank. Sequences type Gene Ascension number Candidatus Borrelia africana TCI22 flaB KT364343 Candidatus Borrelia ivorensis TCI140 flaB KT364344 Candidatus Borrelia ivorensis TCI351 flaB KT364346 Borrelia sp. genotype TCI301 flaB KT364345 Candidatus Borrelia africana TCI22 16S rRNA KT364339 Candidatus Borrelia ivorensis TCI140 16S rRNA KT364340 Candidatus Borrelia ivorensis TCI351 16S rRNA KT364341 Borrelia sp. TCI301 16S rRNA KT364342 Candidatus Anaplasma ivorensis TCI50 23S rRNA KT364326 Candidatus Anaplasma ivorensis TCI94 23S rRNA KT364327 Candidatus Anaplasma ivorensis TCI149 23S rRNA KT364328 Candidatus Wolbachia ivorensis TCI113 23S rRNA KT364329 Candidatus Ehrlichia rustica TCI141 23S rRNA KT364330 Candidatus Ehrlichia rustica TCI145 23S rRNA KT364331 Candidatus Ehrlichia rustica TCI167 23S rRNA KT364332 Candidatus Ehrlichia rustica TCI238 23S rRNA KT364333 Candidatus Ehrlichia urmitei TCI148 23Sr RNA KT364334 Candidatus Ehrlichia urmitei TCI230 23S rRNA KT364335 Candidatus Ehrlichia urmitei TCI106 23S rRNA KT364336 Candidatus Ehrlichia urmitei TCI127 23S rRNA KT364337 Candidatus Ehrlichia urmitei TCI166 23S rRNA KT364338 doi:10.1371/journal.pntd.0004367.t004 correspond to new species. After the construction of a phylogenetic tree (Fig 3), we propose that the status of Candidatus is applied to an uncultured species but not formally recognized by the International Code of Nomenclature of Bacteria [62]. The result shows three cases of Anaplasma: Candidatus Anaplasma ivorensis related to A. phagocytophilum identified in ticks, two in Am. variegatum, and one in Rh. microplus. The three sequences have one to two SNP (single nucleo- tide polymorphism) between them. In one Rh. microplus, a potential new Wolbachia sp., Candi- datus Wolbachia ivorensis, was identified, closely related to the Wolbachia endosymbiont of Cimex lectularius (GenBank AP013028). We also identified two groups of sequences correspond- ing to new Ehrlichia spp. which cluster in two clades. Indeed, in four cases (one Am. variegatum, two Rh.microplus,andoneH. truncatum), we identified Candidatus Ehrlichia rustica in the sub- group of Ehrlichia chaffeensis. In nine ticks (five Am. variegatum,threeRh. microplus and one H. truncatum), we detected Candidatus Ehrlichia urmitei that was previously observed by our team in Rh. bursa ticks collected in the Bacque area of France (M. Dahmani, personal communication) (Fig 3). Candidatus Ehrlichia urmitei forms an independent and well-supported clade situated between the E. ruminantium clade and that of Ehrlichia muris (Fig 3). Finally, 15 co-infections (15/378; 4%) were detected by qPCR. All 15 co-infections involved the presence of R. africae.InAm. variegatum, ten co-infections (10/15; 66%) were observed with R. africae plus another pathogen such as Coxiella burnetii (1/170; 0.6%), A. centrale (1/170; 0.6%), A. marginale (1/170; 0.6%), Candidatus Borrelia Africana, Candidatus Borrelia ivorensis (3/170; 2%), Candidatus Anaplasma ivorensis (2/170; 1%), or Candidatus Ehrlichia urmitei (2/170; 1%) as well as H. marginatum rufipes with R. africae plus R. aeschlimannii (2/ 16; 12%) and in Rh. senegalensis with R. africae plus R. aeschlimannii (1/3; 33%) (Table 3). The access numbers of the sequences of all the potential new species deposited in GenBank are summarized in Table 4. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 12 / 18 264 / 285 Pathogens in Tick Vectors in Côte d'Ivoire Discussion Domestic animal resources supply some 30% of total human food and agricultural production requirements. They are particularly vital to subsistence and economic development in develop- ing countries as they continually provide essential food products, draught power and manure for crop production and generate income as well as employment for most of the rural poor [63]. However, livestock-associated ticks are often reservoirs or vectors of human vector-borne diseases [18]. Intensification of livestock farming is one cause of the abundance of various vec- tors and tick-borne diseases. In recent years, the spectrum of tick-borne diseases infecting ani- mals has increased; many of these diseases, such as rickettsioses, borrelioses, Q fever, anaplasmoses, and ehrlichioses, are gaining increasing attention from clinicians and veterinar- ian [4]. Advances in the development of molecular biology tools facilitate the detection of new bacteria [4,64]. Rickettsioses have been identified in humans, animals and ticks which are considered to be the main vectors of such pathogens as R. africae, R. aeschlimannii, and R. massiliae in sub- Saharan Africa [9]. In our study, rickettsial DNA was found in 49% of ticks collected from cat- tle. For the first time, the presence of R. aeschlimannii in ticks in Côte d’Ivoire is shown. This study provides evidence of R. aeschlimannii infection in 23% of Hyalomma ticks including H. marginatum rufipes (44%) and H. truncatum (11%). R. aeschlimannii has not been observed in other tick species. These data support the theory that the Hyalomma genus is a main vector and reservoir of R. aeschlimannii. It was previously reported in 45% to 51% of H. marginatum rufipes and 6% to 7% in H. truncatum collected from cows, donkeys, sheep, goats and horses in Senegal [12]. These data are comparable to those of our study. The high prevalence of R. africae (90%) in Am. variegatum can be explained by the high transovarial and trans-stadial transmis- sion rates (100%) and a filial infection rate (93%) that was previously demonstrated in Am. var- iegatum [20]. This result shows that this tick species acts as a vector but also as a reservoir for R. africae in Côte d’Ivoire. R. africae was recently detected in other tick genera including Rhipi- cephalus and Hyalomma [12,18,19,57]. In our study, the prevalence of R. africae is 10% in Rh. microplus and 9% in Hyalomma spp., which is lower than in co-fed Am. variegatum, suggesting that these ticks are probably not the competent vectors for R. africae. This bacterium likely infects Rh. microplus and Hyalomma spp. during co-feeding. The first report of the presence of R. massiliae in Côte d’Ivoire was in Rhipicephalus spp. [28]; this is comparable to the detection of R. massiliae in a Rh. senegalensis tick found in our study. C. burnetii infections have been also reported as being between 0.7% and 6.8% in ticks from cattle in western African countries [51] but not in Côte d’Ivoire where the seroprevalence of C. burnetii was estimated to be 3% [52]. Here, we show for the first time the presence of C. burne- tii in Côte d’Ivoire, although only in one tick. Most Borrelia species such as B. hispanica, B. dut- tonii, and B. crocidurae detected in Africa, are related to soft ticks. Their main vectors are Ornithodoros spp. [65]. To date, Borrelia sp. was identified only once in an African hard tick, Am. cohaerens, in Ethiopia [39]. It has been also reported that Rhipicephalus spp. transmits Borrelia theileri to cattle, causing bovine borreliosis [18]. In Côte d’Ivoire, we show that Am. variegatum were infected by three potential new Borrelia and Rh. microplus by one potential new Borrelia. The sequences of Borrelia sp. (genotype TCI301) were identical to 99% of those of Borrelia sp. found in engorged Rhipicephalus sp. ticks collected from horse in Brazil (EF141022). Phylogenetic analysis showed that Borrelia sp. TCI301 is classified in the relapsing fever group, close to B. crocidurae and B. hispanica, two etiological agents of relapsing fever in Africa [26,31]. Blast analysis of the flaB gene showed that three new Candidatus Borrelia ivor- ensis and Candidatus Borrelia africana borreliae were significantly different to all other borre- liae, except for this Borrelia sp in Am. cohaerens in Ethiopia [39]. These potential new Borrelia PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 13 / 18 265 / 285 Pathogens in Tick Vectors in Côte d'Ivoire form a new clade between the clades of Lyme disease borreliae and relapsing fever borreliae. Thus, this is the first time that Borrelia species have been detected in Côte d’Ivoire and the first time their presence has been confirmed in hard ticks in Africa. Bacteria from the Anaplasmataceae family were previously known to be pathogens of veteri- nary importance. However, in the three last decades, many human pathogens have been identi- fied in this family [66]. Recently, based on the rrl gene, our team developed new tools to identify most bacteria belonging to Anaplasmataceae family [47]. These tools combine a qPCR followed by a standard PCR then sequencing, and have been used successfully to amplify DNA from bacteria belonging to Anaplasma spp., Ehrlichia spp., Neorickettsia spp., and Wolbachia spp. available in our laboratory [67]. We have successfully amplified Anaplasmataceae DNA in 6% of our ticks. For the first time, we have demonstrated the presence of A. marginale, A. cen- trale, E. ruminantium, and potential novel Ehrlichia, Anaplasma, and Wolbachia spp. in ticks in Côte d’Ivoire. A. marginale was observed in Am. variegatum and Rh. microplus. To the best of our knowledge, A. marginale has never been reported in Africa in Rh. microplus. The first report of its presence in Côte d’Ivoire was in 2007, but the exact route of its introduction into this region has not yet been determined [68]. A recent study indicated that the majority of the Rhipicephalus (ex-Boophilus) spp. collected and identified from farms around Azaguié (Côte d’Ivoire) are Rh. microplus (96%) [69]. A. centrale is a species closely related to A. marginale; this naturally attenuated strain has been used as a live vaccine to prevent severe diseases due to A. marginale senso stricto strains for 100 years [70]. We identified this species in Am. variega- tum. To the best of our knowledge, A. centrale has never previously been detected in these ticks. The potential of Am. variegatum to transmit A. centrale needs further investigation. E. ruminantium was previously described in Am. variegatum which is invasive to cattle attaches to the hooves and cattle remain standing, particularly in the rainy season [71,72]. Recent phylo- genetic analyses of Am. variegatum from Kenya, Mali, Burkina Faso, Ethiopia and the Carib- bean show low genetic diversity within this population, suggesting an westward expansion of these ticks and supporting east-west genetic separation, with Caribbean genetic sequences being associated with and often identical to West African haplotypes. The data suggest that Am. variegatum reached West Africa from Zambia [73]. We have also identified three potential new species, Candidatus Anaplasma ivorensis, Candidatus Ehrlichia urmitei, and Candidatus Ehrlichia rustica. The detection of these potential new species has limitations, as not all previ- ously described species are already molecularly characterized. Indeed, such species as Ana- plasma caudatum, Anaplasma bovis, and Anaplasma mesaeterum [74] are incompletely characterized with no strain available and no or few genes sequenced, so the detection of a ‘new’ genotype may, in fact, be the re-discovery of an old, incompletely characterized species. Further studies are required to clarify whether these new genetic variants represent a new spe- cies. The other potential new ehrlichiae was closely related to Ehrlichia sp. amplified from Rh. bursa in France. Interestingly, this new species was amplified from two different regions in the world and from different species of ticks (Rhipicephalus, Amblyomma, and Hyalomma spp). Finally, it is reported that ticks are often co-infected following a blood meal from a co- infected host [75,76]. Recently, mixed infections were reported for the first time in West Africa in feeding ticks and caused mainly by Rickettsia spp. and C. burnetii [18]. In Côte d’Ivoire, for the first time we show multiple co-infections in ticks. These co-infections systematically involved R. africae. To date, no human cases of anaplasmoses, ehrlichioses, borrelioses, rickett- sioses or co-infections have been reported in Côte d’Ivoire. However, these diseases are still lit- tle known by clinicians and laboratory diagnostic is lacking in most cases. It is important to continue to study the epidemiological data of such emerging pathogens which may be the source of disease complications in both animals and humans. We provide evidence and dem- onstrate the endemicity of these different bacteria in the studied regions that have the same PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 14 / 18 266 / 285 Pathogens in Tick Vectors in Côte d'Ivoire characteristics agro-ecological and climatic. Furthermore, these diseases could be a cause of death of unknown origin in rural areas in Côte d'Ivoire [77]. Acknowledgments We thank Ange Goun (IRD, Abidjan) for his help in logistic issues. Author Contributions Conceived and designed the experiments: KPY FF OM. Performed the experiments: CBE KPY MD YLA NA AKN JDN OM. Analyzed the data: CBE KPY MD DR FF OM. Contributed reagents/materials/analysis tools: CBE KPY YLA AKN JDN. Wrote the paper: CBE DR FF OM. References 1. Bechah Y, Socolovschi C, Raoult D. Identification of rickettsial infections by using cutaneous swab specimens and PCR. Emerg Infect Dis. 2011; 17(1):83–6. doi: 10.3201/eid1701.100854 PMID: 21192860. 2. Walker AR, Bouattour A, Camicas J, Estrada-Pena A, Horak IG, Latif AA, et al. Ticks of Domestic Ani- mals in Africa: A Guide to Identification of Species. Published by Bioscience Reports, Edinburgh, United Kingdom. 2003. http://www.biosciencereports.pwp.blueyonder.co.uk/. 3. Pérez-Eid C. Emergence of tick-borne diseases in temperate countries. Ann Biol Clin (Paris). 2004; 62 (2):149–54. PMID: 15047466. 4. Dantas-Torres F, Chomel BB, Otranto D. Ticks and tick-borne diseases: a One Health perspective. Trends Parasitol. 2012; 28(10):437–46. doi: 10.1016/j.pt.2012.07.003 PMID: 22902521. 5. Robyn MP, Newman AP, Amato M, Walawander M, Kothe C, Nerone JD, et al. Q Fever Outbreak Among Travelers to Germany Who Received Live Cell Therapy -United States and Canada, 2014. MMWR Morb Mortal Wkly Rep. 2015, 2; 64(38):107. doi: 10.15585/mmwr.mm6438a3 PMID: 26421460. 6. Sokhna C, Mediannikov O, Fenollar F, Bassene H, Diatta G, Tall A, et al. Point-of-care laboratory of pathogen diagnosis in rural Senegal. PLoS Negl Trop Dis. 2013; 7(1):e1999. doi: 10.1371/journal.pntd. 0001999 PMID: 23350001. 7. Mayxay M, Sengvilaipaseuth O, Chanthongthip A, Dubot-Pérès A, Rolain JM, Parola P, et al. Causes of Fever in Rural Southern Laos. Am J Trop Med Hyg. 2015; 93(3):517–20. doi: 10.4269/ajtmh.14- 0772 PMID: 26149859. 8. Ghosh L, Nagar G. Problem of tics and tick-borne diseases in India with a special emphasis on prog- ress in tick control research: a review. J Vector Borne. 2014; 51 (4): 259–70. PMID: 25540956. 9. Parola P, Paddock CD, Socolovschi C, Labruna MB, Mediannikov O, Kernif T, et al. Update on tick- borne rickettsioses around the world: a geographic approach. Clin Microbiol Rev. 2013; 26 (4):657– 702. doi: 10.1128/CMR.00032-13 PMID: 24092850. 10. Kelly PJ, Beati L, Mason PR, Matthewman LA, Roux, Raoult D, et al. Rickettsia africae sp. nov., the etio- logical agent of African tick bite fever. Int J Syst Bacteriol.1996; 46 (2):611–4. PMID: 8934912. 11. Ndip LM, Fokam EB, Bouyer DH, Ndip RN, Titanji VP, Walker DH, et al. Detection of Rickettsia africae in patients and ticks along the coastal region of Cameroon. Am J Trop Med Hyg. 2004; 71:363–6. PMID: 15381820. 12. Mediannikov O, Diatta G, Fenollar F, Sokhna C, Trape JF, Raoult D. Tick-borne rickettsioses, neglected emerging diseases in rural Senegal. PLoS Negl Trop Dis. 2010; 14; 4(9). pii: e821. doi: 10. 1371/journal.pntd.0000821 PMID: 20856858. 13. Parola P, Inokuma H, Camicas JL, Brouqui P, Raoult D. Detection and identification of spotted fever group Rickettsiae and Ehrlichiae in African ticks. Emerg Infect Dis. 2001; 7(6):1014–7. PMID: 11747731. 14. Parola P. Rickettsioses in sub-Saharan Africa. Ann N Y Acad Sci. 2006; 1078:42–7. PMID: 17114679. 15. Socolovschi C, Matsumoto K, Marie JL, Davoust B, Raoult D, Parola P. Identification of Rickettsiae, Uganda and Djibouti. Emerg Infect Dis. 2007; 13 (10):1508–10. doi: 10.3201/eid1310.070078 PMID: 18257998. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 15 / 18 267 / 285 Pathogens in Tick Vectors in Côte d'Ivoire 16. Mediannikov O, Davoust B, Socolovschi C, Tshilolo L, Raoult D, Parola P. Spotted fever group rickett- siae in ticks and fleas from the Democratic Republic of the Congo. Ticks Tick Borne Dis. 2012; 3(5– 6):371–3. doi: 10.1016/j.ttbdis.2012.10.015 PMID: 23137572. 17. Mediannikov O, Diatta G, Zolia Y, Balde MC, Kohar H, Trape JF, et al. Tick-borne rickettsiae in Guinea and Liberia. Tick Borne Dis. 2012; 3(1):43–8. doi: 10.1016/j.ttbdis.2011.08.002 PMID: 22309858. 18. Reye AL, Arinola OG, Hubschen JM, Muller CP. Pathogen prevalence in ticks collected from the vege- tation and livestock in Nigeria. Appl Environ Microbiol. 2012; 78(8):2562–8. doi: 10.1128/AEM.06686- 11 PMID: 22327584. 19. Ogo NI, de Mera IG, Galindo RC, Okubanjo OO, Inuwa HM, Agbede RI, et al. Molecular identification of tick-borne pathogens in Nigerian ticks. Vet Parasitol. 2012; 187(3–4):572–7. doi: 10.1016/j.vetpar. 2012.01.029 PMID: 22326937. 20. Socolovschi C, Huynh TP, Davoust B, Gomez J, RaoulT D, Parola P. Transovarial and trans-stadial transmission of Rickettsiae africae in Amblyomma variegatum ticks. Clin Microbiol Infect. 2009; 15 Suppl 2:317–8. doi: 10.1111/j.1460691.2008.02278.x PMID: 19456811. 21. Raoult D, Fournier PE, Abboud P, Caron F. First documented human Rickettsia aeschlimannii infection. Emerg Infect Dis. 2002; 8:748–9. PMID: 12095451. 22. Parola P, Paddock CD., Raoult D. Tick-borne rickettsioses around the world: emerging diseases chal- lenging old concepts. Clin Microbiol Rev.2005, 18(4): 719–56. PMID: 16223955. 23. Kernif T, Djerbouh A, Mediannikov O, Ayach B, Rolain JM, Raoult D, et al. Rickettsia africae in Hya- lomma dromedarii ticks from sub-Saharan Algeria. Ticks Tick Borne Dis. 2012; 3(5–6):377–9. doi: 10. 1016/j.ttbdis.2012.10.013 PMID: 23164496. 24. Kamani J, Baneth G, Apanaskevich DA, Mumcuoglu KY, Harrus S. Molecular detection of Rickettsia aeschlimannii in Hyalomma spp. ticks from camels (Camelus dromedarius) in Nigeria, West Africa. Med Vet Entomol. 2015; 29(2):205–9. doi: 10.1111/mve.12094 PMID: 25565180. 25. Parola P, Socolovschi C, Jeanjean L, Bitam I, Fournier PE, Sotto A, et al. Warmer weather linked to tick attack and emergence of severe rickettsioses. PLoS Negl Trop Dis. 2008; 2(11):e338. doi: 10.1371/ journal.pntd.0000338 PMID: 19015724. 26. Garcia-Garcia JC, Portillo A, Nunez MJ, Santibanez S, Castro B, Oteo JA. A patient from Argentina infected with Rickettsia massiliae. Am J Trop Med Hyg. 2010; 82(4):691–2. doi: 10.4269/ajtmh.2010. 09–0662 PMID: 20348520. 27. Cascio A, Torina A, Valenzise M, Blanda V, Camarda N, Bombaci S, et al. Scalp escharand neck lymphadenopathy caused by Rickettsia massiliae. Emerg Infect Dis. 2013; 19(5):836–7. doi: 10.3201/ eid1905.121169 PMID: 23697545. 28. Berrelha J, Briolant S, Muller F, Rolain JM, Marie JL, Pagés F, et al. Rickettsia felis and Rickettsia mas- siliae in Ivory Coast, Africa. Clin Microbiol Infect. 2009; 15 Suppl 2:251–2. doi: 10.1111/j.1469-0691. 2008.02273.x PMID: 19548990. 29. Barbour AG. Fall and rise of Lyme of disease and other Ixodes tick-borne infections in North America and Europe. Br Med Bull. 1998; 54(3):647–58. PMID: 10326291. 30. Cutler SJ, Abdissa A, Trape JF. New concepts for the old challenge of African relapsing fever borrelio- sis. Clin Microbiol Infect. 2009; 15(5):400–6. doi: 10.1111/j.1469-0691.2009.02819.x PMID: 19489922. 31. Trape JF, Diatta G, Arnathau C, Bitam I, Sarih M, Belghyti D, et al. The epidemiology and geographic distribution of relapsing fever borreliosis in West and North Africa, with a review of the Ornithodoros erraticus complex (Acari: Ixodida). PLoS One. 2013; 8(11):e78473. doi: 10.1371/journal.pone. 0078473 eCollection2013. PMID: 24223812. 32. Svihrova V, Hudeckova H, Jesenak M, Schwarzova K, Kostanova Z, Ciznar I, et al. Lyme borreliosis- analysis of the trends in Slovakia, 1999–2008. Folia Microbiol (Praha). 2011; 56(3):270–5. doi: 10. 1007/s12223-011-0036-y PMID: 21607749. 33. Vial L, Diatta G, Tall A, Ba el H, Bouganali H, Durand P, et al. Incidence of tick-borne relapsing fever in West Africa: longitudinal study. Lancet. 2006; 368(9529):37–43. PMID: 16815378. 34. Parola P, Ryelandt J, Mangold AJ, Mediannikov O, Guglielmone AA, Raoult D, et al. Relapsing fever Borrelia in Ornithodoros ticks from Bolivia. Ann Trop Med Parasitol. 2011; 105(5):407–11. doi: 10. 1179/1364859411Y.0000000021 PMID: 21929883. 35. Cutler SJ, Bonilla EM, Singh RJ. Population structure of East African relapsing fever Borrelia spp. Emerg Infect Dis. 2010; 16(7):1076–80. doi: 10.3201/eid1607.091085 PMID: 20587177. 36. Sarih M M Garnier, Boudebouch N, Bouattour A, Rihani A, Hassar M, et al. Borrelia hispanica relapsing fever, Morocco. Emerg Infect Dis. 10: 1626–1629. Emerg Infect Dis. 2009; 15(10):1626–9. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 16 / 18 268 / 285 Pathogens in Tick Vectors in Côte d'Ivoire 37. Trape JF, Godeluck B, Diatta G, Rogier C, Legros F, Albergel J, et al. The spread of tick-borne borrelio- sis in West Africa and its relationship to sub-Saharan drought. Am J Trop Med Hyg. 1996; 54(3):289– 93. PMID: 8600768. 38. Nordstrand A, Bunikis I, Larsson C, Tsogbe K, Schwan TG, Nilsson M, et al.Tickborne relapsing fever diagnosis obscured by malaria, Togo. Emerg Infect Dis. 2007; 13(1):117–23. PMID: 17370524. 39. Mediannikov O, Abdissa A, Socolovschi C, Diatta G, Trape JF, Raoult D, et al. Detection of a new Bor- relia species in ticks taken from cattle in Southwest Ethiopia. Vector Borne Zoonotic Dis. 2013; 13 (4):266–9. doi: 10.1089/vbz.2011.0874 PMID: 23421894. 40. Pruneau L, Moumène A, Meyer DF, Marcelino I, Lefrançois T, Vachiéry N, et al. Understanding Ana- plasmataceae pathogenesis using “Omics” approaches. Front Cell Infect Microbiol. 2014; 4:86. doi: 10. 3389/fcimb.2014.00086. eCollection 2014 PMID: 25072029. 41. Rymaszewska. A, Grenda S. Bacteria of the genus Anaplasma–characteristics of Anaplasma and their vectors: a review. Veterinarni Medicina. 2008. 53 (11): 573–584. 42. Aubry P, Geale DW. A review of bovine anaplasmosis. Transbound Emerg Dis. 2011; 58(1):1–30. doi: 10.1111/j.1865-1682.2010.01173.x PMID: 21040509. 43. Raoult D, Parola P. Molecular tools in the epidemiology of tick–borne bacterial diseases Ann Biol Clin (Paris). 2001. 59(2):177–82. PMID: 11282521. 44. Georges M, Garrity. Bergey’s of Manual of Systematic Bacteriology: The proteobacteria. Second Edi- tion. Volume two; 2005. pp.120-122. 45. Allsopp BA. Natural history of Ehrlichia ruminantium. Vet Parasitol. 2010; 167(2–4):123–35. doi: 10. 1016/j.vetpar.2009.09.014 PMID: 19836892. 46. Djiba ML, Mediannikov O, Mbengue M, Thiongane Y, Molez JF, Seck MT, et al. Survey of Anaplasma- taceae bacteria in sheep from Senegal. Trop Anim Health Prod. 2013; 45(7):1557–61. doi: 10.1007/ s11250-013-0399-y PMID: 23553260. 47. Dahmani M, Bernard D, Mohamed SB, Fenollar F, Raoult D, Mediannikov O. Development of a new PCR-based assay to detect Anaplasmataceae and the first report of Anaplasma phagocytophilum and Anaplasma platys in cattle from Algeria. Comp Immunol Microbiol Infect Dis. 2015; 39:39–45. doi: 10. 1016/j.cimid.2015.02.002 PMID: 25748051. 48. Maurin M, Raoult D. Q fever. Clin Microbiol Rev. 1999; 12(4):518–53. PMID: 10515901. 49. Kazar J. Coxiella burnetii infection. Ann N Y Acad Sci. 2005; 1063:105–14. PMID: 16481501. 50. Cooper A, Hedlefs R, Ketheesan N, Govan B. Serological evidence of Coxiella burnetii infection in dogs in a regional centre. Aust Vet J. 2011; 89(10):385–7. doi: 10.1111/j.1751-0813.2011.00819.x PMID: 21933165. 51. Mediannikov O. Coxiella burnetii in humans and ticks in rural Senegal. PLoS Negl Trop Dis. 2010; 4(4): e654. doi: 10.1371/journal.pntd.0000654 PMID: 20386603. 52. Tissot-Dupont H, Brouqui P, Faugere B, Raoult D. Prevalence of antibodies to Coxiella burnetii, Rick- ettsia conorii, and Rickettsia typhi in seven African countries. Clin Infect Dis. 1995; 21(5):1126–33. PMID: 8589132. 53. Rolain JM, Sthul L, Maurin M, Raoult D. Evaluation of antibiotic susceptibilities of three rickettsial spe- cies including Rickettsia felis by a quantitative PCR DNA assay. Antimicrob Agents Chemother. 2002; 46(9):2747–51. PMID: 12183224. 54. Socolovschi C, Frederic P, Raoult D. Rickettsia felis in Aedes albopictus mosquitoes, Libreville, Gabon. Emerg Infect Dis. 2012; 18(10):1687–9. doi: 10.3201/eid1810.120178 PMID: 23017437. 55. Subramanian G, Sekevova Z, Raoult D, Medianikov O. Multiple tick-associated bacteria in Ixodes rici- nus from Slovakia. Ticks Tick Borne Dis. 2012; 3(5–6):406–10. doi: 10.1016/j.ttbdis.2012.10.001 PMID: 23182274. 56. Rolain JM, Raoult D. Molecular detection of Coxiella burnetii in blood and sera during Q fever. QJM. 2005; 98(8):615–7; author reply 617–20. PMID: 16027172. 57. Mediannikov O, Trape JF, Diatta G, Parola P, Fournier PE, Raoult D. Rickettsia africae, Western Africa. Emerg Infect Dis. 2010; 16(3):571–3. doi: 10.3201/eid1603.090346 PMID: 20202453. 58. Raoult D, Roblot F, Rolain JM, Besnier JM, Loulergue J, Bastides, et al. First isolation of Bartonella alsatica from a valve of a patient with endocarditis. J Clin Microbiol. 2006; 44(1):278–9. PMID: 16390990. 59. Bottieau E, Verbruggen E, Aubry C, Socolovschi C, Vlieghe E. Meningoencephalitis complicating relapsing fever in traveler returning from Senegal. Emerg Infect Dis. 2012; 18(4):697–8. doi: 10.3201/ eid1804.111771 PMID: 22469185. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 17 / 18 269 / 285 Pathogens in Tick Vectors in Côte d'Ivoire 60. Mura A, Socolovschi C, Ginesta J, Lafrance B, Magnan S, Rolain JM, et al. Molecular detection of spot- ted fever group rickettsiae in ticks from Ethipia and Chad. Trans R Soc Trop Med Hyg. 2008; 102 (9):945–9. doi: 10.1016/j.trstmh.2008.03.015 PMID: 18440576. 61. Ronquist F, Huelsenbeck JP. MrBayes 3: Bayesian phylogenetic inference under mixed models. Bioin- formatics. 2003; 19 (12):1572–4. PMID: 12912839. 62. Murray RG, Stackebrandt E. Taxonomic note: implementation of the provisional status Candidatus for incompletely described procaryotes. Int J Syst Bacteriol. 1995; 45 (1):186–7 I. PMID: 7857801. 63. Ayalew W, Rege JEO, Getahun E, Tibbo M, Mamo Y. Delivering Systematic Information on Indigenous Animal Genetic Resources–the development and prospects of DAGRIS. Animal Genetic Resources Group, International Livestock Research Institute. P, O Box 5689. 2003. Addis Ababa, Ethiopia. 64. Doudier B, Olano J, Parola P, Brouqui P. Factors contributing to emergence of Ehrlichia and Anaplasma spp. as human pathogens. Vet Parasitol. 2010; 167(2–4):149–54. doi: 10.1016/j.vetpar.2009.09.016 PMID: 19836890. 65. Parola P, Raoult D. Ticks and tick-borne bacterial diseases in humans: an emerging infectious threat. Clin Infect Dis. 2001; 32(6):897–928. PMID: 11247714. 66. Inokuma H. Vectors and reservoir hosts of Anaplasmataceae. Rickettsial diseases, books.google. com. 2007, pp. 199–212. 67. Dahmani M, Loudahi A, Mediannikov O, Fenollar F, Raoult D, Davoust B. Molecular detection of Ana- plasma platys and Ehrlichia canis in dogs from Kabylie, Algeria. Ticks Tick Borne Dis. 2015; 6(2):198– 203. doi: 10.1016/j.ttbdis.2014.12.007 PMID: 25583345. 68. Madder M, Thys E, Geysen D, Baudoux C, Horak I. Boophilus microplus ticks found in West Africa. Exp Appl Acarol. 2007; 43(3):233–4. PMID: 17929178. 69. Madder M, Thys E, Achi L, Touré A, De Deken R. Rhipicephalus (Boophilus) microplus: A most suc- cessful invasive tick species in West-Africa. Exp Appl Acarol. 2011; 53(2):139–45. doi: 10.1007/ s10493-010-9390-8 PMID: 20711801. 70. Herndon DR, Palmer GH, Shkap V, Knowles DP, Brayton K. Complete genome sequence of Ana- plasma marginale sub sp centrale. J Bacteriol. 2010; 192(1):379–80. doi: 10.1128/JB.01330-09 PMID: 19854912. 71. Kifle G, Sori T. Molecular Detection of Ehrlichia Species in Amblyomma Ticks Collected from Rumi- nants in Abernosa Ranch, Ethiopia. Global J Mol Sci. 2014; 9 (2): 12–18. doi: 10.5829/idosi.gjms.2014. 9.2.85213 72. Stachurski F. Invasion of West African cattle by the tick Amblyomma variegatum. Med Vet Entomol. 2000; 14(4):391–9. PMID: 11129703. 73. Beati L, Patel J, Lucas-Williams H, Adakal H, Kanduma EG, Tembo-Mwase E, et al. Phylogeography and demographic history of Ambryomma variegatum (Fabricius) (Acari: Ixodidae), the Tropical Bont Tick. Vector Borne Zoonotic Dis. 2012; 12(6):514–25. doi: 10.1089/vbz.2011.0859 PMID: 22448720. 74. Georges M, Garrity. Bergey’s of Manual of Systematic Bacteriology: The proteobacteria. Second Edi- tion. Volume two; 2005. pp. 121–125. 75. Levin M L, Fish D. Acquisition of coinfection and simultaneous transmission of Borrelia burgdorferi and Ehrlichia phagocytophila by Ixodes scapularis ticks. Infect Immun. 2000; 68(4):2183–6. PMID: 10722618. 76. Levin M L, Fish D. Interference between the agents of Lyme disease and human granulocytic ehrlichio- sis in a natural reservoir host. Vector Borne Zoonotic Dis. 2001; 1(2):139–48. PMID: 12653144. 77. Koné S, Fürst T, Jaeger FN, Esso EL, Baïkoro N, Kouadio KA, et al. Causes of death in the Taabo health and demographic surveillance system, Côte d'Ivoire, from 2009 to 2011. Glob Health Action. 2015; 8:27271. doi: 10.3402/gha.v8.27271. eCollection2015 PMID: 25959772. PLOS Neglected Tropical Diseases | DOI:10.1371/journal.pntd.0004367 January 15, 2016 18 / 18 270 / 285 Article 12 : Molecular identification of microorganisms in chronic wounds, Republic of Guinea (Conakry) Publié dans Global Advanced Research Journal of Microbiology 2018; Vol. 7(1): pp. 023-036 271 / 285 Global Advanced Research Journal of Microbiology (ISSN: 2315-5116) Vol. 7(1) pp. 023-036, February, 2018 Issue. Available online http://garj.org/garjm Copyright© 2018 Global Advanced Research Journals Full Length Research Paper Molecular identification of microorganisms in chronic wounds, Republic of Guinea (Conakry) Cyrille Bilé Ehounoud1,2, Alpha Kabinet Keita3, Abdoul Habib Béavogui4,5, Ansoumane Cissé4, Nadia Amanzougaghene1, Jean David N’Guessan2, Didier Raoult1, Oleg Mediannikov1, Florence Fenollar1* 1Aix Marseille Univ, Unité de Recherche sur les Maladies Infectieuses et Tropicales Emergentes (URMITE), UM63, CNRS 7278, IRD 198, INSERM 1095, IHU - Méditerranée Infection, 19-21 Boulevard Jean Moulin, 13005 Marseille 2Felix Houphouet Boigny Université, UFR Biosciences, Côte D’Ivoire. 3Unité de recherche translationnelle appliquée au VIH et les maladies infectieuses (TransVIHMI) IRD UMI 233-INSERM U 1175 Centre IRD France Sud 911, avenue Acropoles, BP 64501 F-34394 Montpellier cedex 5. 4Centre de Formation et de Recherche en Santé Rurale de Mafèrinyah B.P. 2649, Conakry, République de Guinée. 5Faculté de Médecine, Pharmacie et Odonto-Stomatologie, Université Gamal Abdel Nasser de Conakry, République de Guinée. Accepted 13 January, 2018 Skin infections are common in sub-Saharan Africa, including chronic wounds. This study aimed to assess the presence of several microorganisms in skin specimens from patients with chronic wounds and healthy people in Maferinyah (Republic of Guinea). Eighty-four skin samples from the wounds of 20 patients (42 edge swabs and 42 center swabs) and twenty-two skin samples from 11 healthy people were analyzed by real-time quantitative PCR and standard PCR assays combined with sequencing. Pseudomonas aeruginosa was the most frequently detected bacterium, which was significantly more prevalent in patients (80%, 16/20) than in healthy people (9%, 1/11; p<0.001), followed by Staphylococcus aureus which was only detected in patients (60%, 12/20; p<0.001). Streptococcus pyogenes was also more frequently detected in patients (30%, 6/20) than in healthy people (9%, 1/11) but the difference was not statistical significant. Rickettsia felis was also detected for the first time in Guinea, in one patient. Finally, species of the genus Acinetobacter were also frequently and exclusively detected in patients (80%, 16/20). Acinetobacter baumannii (2/20, 10%), Acinetobacter nosocomialis (10%), Acinetobacter junii (1/20, 5%), Acinetobacter lwofii (5%), and Acinetobacter guangdongensis (5%), which was detected for the first time in skin, were identified. Acinetobacter junii and Acinetobacter lwofii were observed in different samples from the same patient. For the 11 other patients, polymicrobial infections featuring several species of the genus Acinetobacter were observed. Overall, many different bacteria which may encourage wound enlargement or delayed healing were observed in chronic wounds. Keywords: skin; chronic wound; bacteria; Staphylococcus aureus; Pseudomonas aeruginosa; Acinetobacter spp.; Rickettsia felis; Guinea (Conakry) 272 / 285 024. Glo. Adv. Res. J. Microbiol. INTRODUCTION Chronic wounds represent a common pathology in poor Buruli ulcer is another cause of skin disease associated countries, affecting about 15% of the population in Sub- with tropical and humid areas in Africa where there are Saharan Africa compared to 1% in developed countries slow moving rivers and stagnant water (Wansbrough- (Gulam-Abbas et al., 2002; Gottrup, 2004). Diabetic foot Jones and Phillips, 2006). This chronic skin disease is ulcers, venous leg ulcers, surgical wounds, eschars, burns, caused by infection with Mycobacterium ulcerans leading and bites are regularly observed (Rhoads et al., 2012; to the development of large ulcers (Wansbrough-Jones Mediannikov et al., 2014; Essayagh et al., 2014; Pratt et and Phillips, 2006). Environmental sources of M. ulcerans al., 2016). Chronic wounds have a real impact upon are better characterized but the mode of transmission of morbidity and disability, as patients can live for years with infection is still uncertain (Wansbrough-Jones and Phillips, wounds, but also have an impact upon mortality. In a study 2006). The introduction of rational antibiotic therapy has on diabetic foot ulcers in Tanzania (Sub-Saharan Africa), resulted in improvements in the management of the Gulam-Abbas et al. reported that the mortality rate was up disease (Wansbrough-Jones and Phillips, 2006). to 54% among in-patients with severe ulcerations without In Guinea, little is known about the microorganisms amputation of the affected part.1 Smith also speculate that present in chronic wounds. This study aimed to evaluate in future years, the death rate will increase because of the the prevalence of microorganisms in chonic wounds in high costs of care (Smith, 2004), which may often force Guinea (Conakry), including the use of controls (healthy patients to treat themselves at home or to consult skin), as new strategies have emerged indicating the alternative medicine providers. Studies on foot ulcers possible role of a microorganism as the cause of infection. estimate that amputation rates are 45% and 23.5% and mortality is 38% and 9%, respectively, in western and northern African countries (Sano et al., 1998; Benotmane PATIENTS, MATERIAL AND METHODS et al., 2000). These prevalence rates demonstrate the extent of the problem of chronic wounds in which infections Patients and control group represent a major cause of amputations and deaths (Gomez et al., 2009; Sen et al., 2009; Wolcott et al., This study includes 84 skin samples obtained from 20 2010a). patients with chronic skin wounds. These patients, who live Most wounds infections are caused by bacteria. The in rural areas, have consulted at Primary Health Care most common are Staphylococcus aureus, Streptococcus center of Maferinyah, Republic of Guinea (Conakry) in pyogenes, and Pseudomonas aeruginosa (Gjødsbøl et al., June 2014. At the same time, 22 skin specimens from 11 2006; Rhoads et al., 2012; Mediannikov et al., 2014). The healthy people living in the same area were sampled presence of Acinetobacter spp., including Acinetobacter (Table 1). Samples were collected according the protocol baumannii, is also significant in burn wound infections. previously reported by Mediannikov et al (Mediannikov et One study reported that they were the most prevalent al., 2014). Briefly, the cotton swab (Copan, Brescia, Italy) (22.2%) followed by P. aeruginosa (15.1%), and S. aureus was applied firmly to the center and edge of the wound. at 10.3% (Essayagh et al., 2014). Rickettsia felis, a For negative control samples, the cotton swab was applied bacterium involved in fever in sub-Saharan Africa and Asia to the skin (inner surface of the forearm) of healthy people. was reported in eschars (7.4%) but also in skin from All lesions were photographed. All samples were healthy people (5%) in Senegal (Mediannikov et al., 2014; transferred to the URMITE laboratory (Marseille, France). Socolovschi et al., 2010; Ferdouse et al., 2015; The information gathered on each patient included their Mourembou et al., 2015a). age, sex, the presence of fever (axillary temperature > Molecular testing methods present several advantages, 37.5°C), glycemia (0.75 g/L < normal glycemia < 1.10 g/L), such as the ability to identify fastidious bacteria and dead history, evolution of the wound, and use of river water. bacteria following, for example, antibiotic therapies or when None of the patients had received antibiotic treatment or specimens have been kept in poor transport or storage local antiseptic treatment before sampling. conditions. Thus, although molecular methods cannot replace culture in terms of obtaining isolates and data Ethics Statement about antibiotic susceptibilities, they provide a significant amount of information and enable the description of This study was approved by the ethics committee of bacterial repertoire (Thomsen et al., 2010; Wolcott et al., Guinea, Conakry (agreement number 008/CNERS/14). 2010b; Rhoads et al., 2012). Written informed consent from all participants, including patients and the parents or legal guardians of children was obtained. *Corresponding Author’s Email: [email protected] 273 / 285 Ehounoud et al. 025 Table 1. Distribution of samples according to sample area. Sample area Edge Center People Number of samples Total Wounds 20 42 42 84 Healthy skin 11 11 11 22 Molecular analysis extension steps for 30 seconds at 45°C. In each reaction, two positive controls (microbial DNA) and two negative Each cotton swab was put in 200 µL of buffered solution controls (the mix alone) were used to validate each PCR (G2; Qiagen, Hilden, Germany) with 20 µL of proteinase K assay. (Qiagen) and incubated at 56°C for one hour. DNA from For the identification of Acinetobacter species, DNA each sample was extracted using the EZ1 DNA Tissue kit extracts were subjected to standard PCR to amplify a following the manufacturer’s recommendations (Qiagen). portion of the rpoB gene, coupled with sequencing. The The quality of all DNA extracts was checked using primers used (Ac696 Forward quantitative real-time PCR (qPCR) targeting a specific TAYCGYAAAGAYTTGAAAGAAG and Ac1093 Reverse human β-actin gene (primers ActinF 5'- CMACACCYTTGTTMCCRTGA) amplified a 350 bp CATGCCATCCTGCATCTGGA-3' and ActinR 5'- fragment of Acinetobacter rpoB gene, as previously CCGTGGCCATCTCTTGCTCG-3' combined with a reported (La Scola et al., 2006). Standard PCR was TaqMan probe 6-FAM- performed on a ThermalCycler (Applied Biosystem, Paris, CGGGAAATCGTGCGTGACATTAAG-TAMRA), as France). The reactions were carried out using the Hotstar previously reported (Keita et al., 2015; Mourembou et al., Taq-polymerase (Qiagen), in accordance with the 2015b). manufacturer’s instructions. The amplicons were visualized Pathogen screening of skin samples was performed with using electrophoresis on a 1.5% agarose gel stained with qPCR assays using primers and probes targeting S. ethidium bromide and examined using an ultraviolet aureus, S. pyogenes, Streptococcus pneumoniae, transilluminator. The PCR products were purified using a Salmonella spp., Acinetobacter spp., P. aeruginosa, PCR filter plate Millipore NucleoFast 96 PCR kit following Tropheryma whipplei, Rickettsia spp., R. felis, the manufacturer’s recommendations (Macherey-Nagel, Mycobacterium spp., Mycobacterium ulcerans, Düren, Germany) (Ehounoud et al., 2016). The amplicons Mycobacterium marinum, Coxiella burnetii, Treponema were sequenced using the BigDye Terminator Cycle pallidum, Haemophilus ducreyi, Leishmania spp., Sequencing Kit (Applied Biosystems) with an ABI Mansonella spp., and Pox Virus (Table 2) (Rolain et al., automated sequencer (Applied Biosystems). The obtained 2002; Rolain et al., 2005; Leslie et al., 2007; Mediannikov sequences were assembled using ChromasPro software et al., 2010; Bouvresse et al., 2011; Guitard et al., 2012; (ChromasPro 1.7, Technelysium Pty Ltd and Tewantin, Lavender et al., 2012; Hamad et al., 2015; Mourembou et Australia) and compared with those available in GenBank al., 2015a; Mourembou et al., 2015b; Mourembou et al., by NCBI BLAST (http://blast.ncbi.nlm.nih.gov/Blast.cgi) 2016). (Ehounoud et al., 2016). Each PCR assay was performed with a 20 µL volume A phylogenetic tree was constructed by using the test containing 10 µL Master mix No-ROX (Eurogentec, Liege, maximum likelihood in the MEGA6 program Belgium), 3.5 µL of distilled water (DNAase /RNAase free), (http://megasoftware.net/). The Epi Info version 7 program 2.5 µM of probe, 20 µM of each primer, and 5 µL of DNA (http://www.cdc.gov/epiinfo/index.html) was used for data extract (Keita et al., 2015; Mourembou et al, 2015b). All analysis. A difference was statistically significant when p- reactions were performed using a CFX 96 (Bio-Rad, values were <0.05. Marnes-la-Coquette, France) according to the manufacturer’s protocol: DNA denaturation steps at 50°C for two minutes and 95°C for five minutes followed by 40 one-second cycles at 95°C, 35 seconds at 60°C, and 274 / 285 026. Glo. Adv. Res. J. Microbiol. Table 2. Primers and probes used for real-time quantitative PCR in this study. Microorganisms Targeted Primers (5’-3’) References detected sequences Forward Reverse Probes (6 FAM – TAMRA) BACTERIA Coxiella burnetii IS1111 CAAGAAACGTATCGCTGTGGC (Rolain et al., 2005) CACAGAGCCACCGTATGAATC CCGAGTTCGAAACAATGAGGGCTG Hypothetical CGCTGACCTACAGAAATATGTCC (Mediannikov et al., 2010) Protein GGGGTAAGTAAATAATACCTTCTGG CATGAAGCGATTTATCAATACGTGTATGC Haemophilus ducreyi GroESL CACAATGAGTATTCGTCCATTACAC This study GCAATCACTTTACCGCGAGT CGGGTGGTATTGTTTTAACAGGTTCAGCGA Mycobacteria ITS GGGTGGGGTGTGGTGTTTGA (Guitard et al., 2012) CAAGGCATCCACCATGCGC TGGATAGTGGTTGCGAGCATC Mycobacterium ulcerans IS2404 AAAGCACCACGCAGCATCT (Lavender et al., 2012) AGCGACCCCAGTGGATTG CGTCCAACGCGATC Mycobacterium marinum Ppe ATGTGGGCAGCTTCAATGTG This study CCAAGCCAACACTGGAATCA AACATCGGGCCGGGCAACCT Pseudomonas OprL CGCTGCCTTTCAGGTCTTTC This study aeruginosa CGTGCGATCACCACCTTCTA TCCAGAGCGCGCATGGCTTC Hypothetical GAACCGTTGTGCAGGTAGGG This study Protein CGCAAGGACTACTGCCTGAA CGGTGGCCCAGATGCCGTTC Rickettsia spp. RKNDO3 GTGAATGAAAGATTACACTATTTAT (Rolain et al., 2002) GTATCTTAGCAATCATTCTAATAGC CTATTATGCTTGCGGCTGTCGGTTC Rickettsia felis 0527 ATGTTCGGGCTTCCGGTATG (Mourembou et al., 2015a) CCGATTCAGCAGGTTCTTCAA GCTGCGGCGGTATTTTAGGAATGGG OrfB CCCTTTTCGTAACGCTTTGCT (Mourembou et al., 2015a) GGGCTAAACCAGGGAAACCT TGTTCCGGTTTTAACGGCAGATACCCA 275 / 285 Ehounoud et al. 027 Table 2. Continue Salmonella spp. SipC GTCAGGCGTCGTAAAAGCTG (Mourembou et al., 2016) ACGTCGACTGGTGGTACTGG CTCCAGGCGCGAACAGCTGG InvA TCTGTTTACCGGGCATACCA (Mourembou et al., 2016) CACCGTGGTCCAGTTTATCG CCAGAGAAAATCGGGCCGCG Streptococcus PlyN GCGATAGCTTTCTCCAAGTGG (Mourembou et al., 2016) pneumoniae TTAGCCAACAAATCGTTTACCG CCCAGCAATTCAAGTGTTCGCCGA Streptococcus pyogenes Hypothetical ACAGGAACTAATACTGATTGGAAAGG (Mourembou et al., 2016) Protein TGTAAAGTGAAAATAGCAGCTCTAGCA AAAATGTTGTGTTTTAGGCACTGGCGG MipB GGACATAATAAAAGGTTTTTCTTCCA (Mourembou et al., 2016) CAAAATACACAAAATACAGAACCAAA CATTATGATGTGACGTGGTAGGATGGG Staphylococcus aureus NucA TTGATACGCCAGAAACGGTG (Mourembou et al., 2016) TGATGCTTCTTTGCCAAATGG AACCGAATACGCCTGTAC CCTCGACAGGTAACGCATCA (Mourembou et al., 2016) Amidohydrolas AAACTCCTATCGGCCGCAAT e TGCAATGGTAGGTCCTGTGCCCA Tropheryma whipplei TGAGGATGTATCTGTGTATGGGACA (Keita et al., 2015) whi2 TCCTGTTACAAGCAGTACAAAACAAA GAGAGATGGGGTGCAGGACAGGG whi3 TTGTGTATTTGGTATTAGATGAAACAG (Keita et al., 2015) CCCTACAATATGAAACAGCCTTTG GGGATAGAGCAGGAGGTGTCTGTCTGG Acinetobacter spp. rpoB TACTCATATACCGAAAAGAAACGG (Bouvresse et al., 2011) GGYTTACCAAGRCTATACTCAAC CGCGAAGATATCGGTCTSCAAGC Acinetobacter baumannii Nacetyl ARCGGATGCCAAGAGAATGT This study glutamate CCGACATTCAGCACCCTACA synthase GCGGACTGCTTCACCGCCAA pap AAAAAGAGCGTGCACGACAA This study TCGGCCCAAAAATAACTTGG GCGCAAGCGGGTACAACGTGA 276 / 285 028. Glo. Adv. Res. J. Microbiol. Table 2. Continue Treponema pallidum polA GTCGAGACTGAAAAGGAGTGCA (Leslie et al., 2007) GTGAGCGTCTCATCATTCCAAAG TGCTGTGCAGGATCCGGCATATGTCC PARASITES Leishmania spp 18S ACAAGTGCTTTCCCATCG (Hamad et al., 2011) CCTAGAGGCCGTGAGTTG CGGTTCGGTGTGTGGCGCC Mansonella spp ITS CCTGCGGAAGGATCATTAAC (Mourembou et al., 2015b) ATCGACGGTTTAGGCGATAA CGGTGATATTCGTTGGTGTCT VIRUS Pox Virus Hemagglutinin TGATGCAACTCTATCATGTARTCG This study CAAGACGTCGCTTTTRGCAG TGCTTGGTATAAGGAGCCCAATTCCA B2L CGGTGCAGCACGAGGTC This study CGGCGTATTCTTCTCGGACT GCCTAGGAAGCGCTCCGGCG Figure 1. Map of Guinea (Conakry) showing the area of Maferinyah where people were recruited. RESULTS ages ranged from 13 to 90 year-old (mean age 47). Of them, six patients (6/20; 30%) presented a fever and eight Description of the population (8/20; 40%) presented hyperglycemia (glycemia ≥ 1.10 g/L). Eleven patients (11/20; 55%) used the water from Twenty patients (8 males and 12 females) with chronic river to wash their bodies, kitchenware, and linen. wounds, residing in the rural area of Maferinyah, which is Patients had presented wounds from between three crossed by the Kili river, were included (Figure 1). Their weeks and 19 years. Most of the wounds (16/20; 80%) 277 / 285 Ehounoud et al. 029 Table 3. Main epidemiological, biological, and clinical data of patients with chronic wounds. Data Patients Temperature Glycemia History of Evolution of Localization of Use of river water (≥37.5°) (≥1.10) wound wound wound for their needs 1 no no Swelling 2 years External malleolus yes 2 no yes Minor injury NA Back foot yes 3 yes no Trauma object 2 years Back foot no 4 no no Minor injury 3 years External malleolus no 5 no yes Sharp object 10 years Above the ankle yes 6 no no Swelling 5 months Back foot no 7 no no Swelling 4 months External malleolus yes 8 no no Swelling 5 years Back foot yes 9 no no Minor injury 3 weeks Back foot no 10 no yes Snakebite 4 years Back foot yes 11 yes no Wood 7 years Back foot no 12 yes no Snakebite 2 years Above the ankle NA 13 no yes Swelling 19 years Back foot yes 14 yes yes Palm thorn 3 years Back foot yes 15 yes yes Minor injury 8 months External malleolus yes 16 no no Falling tree 1 year Above the ankle yes 17 no no Snakebite 6 years Back foot no 18 no no Sharp object 5 years Sole no 19 no yes NA 6 months Above the ankle no 20 yes yes NA 3 months Foot / Above the yes ankle NA: Not available were located on the foot, such as the back of the foot for taken from patients (66.6%) and in one of the 22 skin 11 patients, external malleolus for four patients, and the samples taken from healthy people (4.5%). P. aeruginosa sole in one case. For the remaining five patients (5/20; was also detected at approximately the same prevalence 25%), wounds were located above the ankle. The wounds at the edges (69%, 29/42) and centers (64.2%, 27/42; had been caused by a sharp object for four patients (4/20; p=0.6) of the wounds. 20%), by a snake bite for three patients (3/20; 15%), by a S. aureus was the second more common microorganism scratch for two patients (2/20; 10%), and by other means identified among patients (60%, 12/20). S. aureus was also for the last 11 patients (Table 3). statistically more frequently detected among patients, as it was not detected in any skin samples taken from healthy Microorganisms detected people (0% versus 60%, p<0.001). In addition, S. aureus was observed in 29 of the 84 skin samples taken from Because ß-actin qPCR was positive for all samples, patients (34.5%). Overall, S. aureus was more frequently revealing the good quality of DNA extracts, all specimens detected at the edges of wounds (40.4%, 17/42) than in the were included in the analyses. All the patients as well as centers (28.5%, 12/42), although the difference was not two of the 11 healthy people were positive for at least one statistically significant (p=0.3). bacterium (Table 4). All the detected bacteria are S. pyogenes was more frequently observed in patients summarized in Table 5. (30%, 6/20) than in healthy people (9%, 1/11) but the P. aeruginosa was the most frequently observed difference was not statistically significant (p=0.1). Overall, microorganism in both patients (80%, 16/20) and healthy the bacterium was found in 15 of the 84 skin samples people (9%, 1/11). However, the bacterium was statistically taken from patients (17.8%) and in one of the 22 healthy more frequently observed among patients than the control skin samples (4.5%) but there was no significant statistical group with healthy skin (80% versus 9%, p<0.001). Overall, P. aeruginosa was observed in 56 of the 84 skin samples 278 / 285 030. Glo. Adv. Res. J. Microbiol. Table 4. Microorganisms identified in skin samples for each patient and healthy people (*Acinetobacter sp P signifies that several species of Acinetobacter were present in the specimens). People Detected microorganisms Center swab Edge swab Total Wounds Patient 1 P. aeruginosa + "Acinetobacter P. aeruginosa P. aeruginosa + "Acinetobacter sp P*" sp P" Patient 2 P. aeruginosa + S. aureus P. aeruginosa P. aeruginosa + S. aureus Patient 3 P. aeruginosa + "Acinetobacter P. aeruginosa + "Acinetobacter P. aeruginosa + "Acinetobacter sp P" spP" + sp P" + S. aureus S. aureus Patient 4 P. aeruginosa P. aeruginosa P. aeruginosa Patient 5 P. aeruginosa + S. aureus P. aeruginosa + S. aureus P. aeruginosa + S. aureus Patient 6 P. aeruginosa + "Acinetobacter P. aeruginosa +"Acinetobacter sp P. aeruginosa + "Acinetobacter 279 /285 sp P" P" + sp P" + S. pyogenes S. pyogenes Patient 7 P. aeruginosa + "Acinetobacter P. aeruginosa + Acinetobacter P. aeruginosa + "Acinetobacter sp P" spp sp P" Patient 8 S. aureus + S. pyogenes S. pyogenes + "Acinetobacter sp S. aureus + S. pyogenes + P" "Acinetobacter sp P" Patient 9 P. aeruginosa + A. baumannii P. aeruginosa + A. baumannii P. aeruginosa + A. baumannii S. aureus + S pyogenes S. aureus + S. pyogenes Patient 10 P. aeruginosa + R. felis + P. aeruginosa + "Acinetobacter sp P. aeruginosa + R. felis + "Acinetobacter sp P" P" "Acinetobacter sp P" Patient 11 S. pyogenes S. aureus + S. pyogenes + S. aureus + S. pyogenes + A. nosocomialis A. nosocomialis Patient 12 P. aeruginosa P. aeruginosa + S. aureus P. aeruginosa + S. aureus Patient 13 P. aeruginosa + S. aureus P. aeruginosa + S. aureus P. aeruginosa + S. aureus + A. nosocomialis + A. nosocomialis Patient 14 A. baumannii A. baumannii A. baumannii Patient 15 P. aeruginosa + "Acinetobacter "Acinetobacter sp P" P. aeruginosa + "Acinetobacter sp P" sp P" Ehounoud et al. 031 Table 4. Continue Patient 16 P. aeruginosa + S. aureus + P. aeruginosa + S. aureus + P. aeruginosa + S. aureus + "Acinetobacter sp P" "Acinetobacter sp P" "Acinetobacter sp P" Patient 17 P. aeruginosa + S. aureus P. aeruginosa + S. aureus P. aeruginosa + S. aureus + A. + A. guangdongensis + A. guangdongensis guangdongensis Patient 18 P. aeruginosa + S. aureus + P. aeruginosa + S. aureus + P. aeruginosa + S. aureus + "Acinetobacter sp P" "Acinetobacter sp P" "Acinetobacter sp P" Patient 19 S. pyogenes + "Acinetobacter sp S. pyogenes + Acinetobacter spp S. pyogenes + "Acinetobacter P" sp P" Patient 20 S. aureus + A. lwoffii P. aeruginosa + S. pyogenes + A. P. aeruginosa + S. aureus + S. junii pyogenes+ A. junii + A. lwoffii Healthy people 1 None P. aeruginosa P. aeruginosa None Healthy people 2 None None None None Healthy people 3 None None None None Healthy people 4 None None None None Healthy people 5 None None None None 280 /285 Healthy people 6 None None None None Healthy people 7 None None None None Healthy people 8 None None None None Healthy people 9 None None None None Healthy people 10 None S. pyogenes S. pyogenes None Healthy people 11 None None None None difference (p=0.1). The edges of wounds were identified. Of all the species of Acinetobacter was observed in 7.1% (6/84) of the specimens: more often tested positive for S. pyogenes identified, A. baumannii was detected in two 9.5% (4/42) from the edges of wounds and 4.7% (23.8%, 10/42) than the centers (11.9%, 5/42) patients (2/20, 10% versus 0% in healthy people; (2/42; p=0.3) from the center. but the difference was not statistically significant p=0.4). For the two patients, the similarity was Acinetobacter guangdongensis was identified (p=0.1). R. felis was detected in only one patient 100% with A. baumannii, previously detected in in one patient (1/20, 5% versus 0% in healthy (5%, 1/20) but not in the skin of healthy people tissue in Germany (Genbank LN868200). people; p=0.6). There was 99% of similarity with (p=0.6). In addition, the bacterium was found in Overall, A. baumannii was observed in seven of the A. guangdongensis strain ANC5077 one sample taken from the edge of the wound the 84 specimens (8.3%). Of them, 9.5% (4/42) (KR611818.1). Overall, A. guangdongensis was (1.1%, 1/84). were detected from the edges of wounds and detected in 4.7% (4/84) of the samples: two from Acinetobacter spp. was only identified in 7.1% (3/42; p=0.5) from the centers. the edges and two from the center of the wound. patients (80%, 16/20) and was not found in the Acinetobacter nosocomialis was detected in skin samples taken from healthy people (0%, two other patients (10%, 2/20 versus 0% in 0/11; p<0.001). Acinetobacter spp. was found in healthy people; p=0.4). There was 100% 47 of the 84 skin samples taken from patients similarity with the A. nosocomialis strain A196 (55.9%). For twenty-one patients (21/47; 44.6%), (KJ788897) for one patient and the LMG10619 the species of Acinetobacter was successfully strain (LC102686) for the other. A. nosocomialis 032. Glo. Adv. Res. J. Microbiol. Table 5. Prevalence of microorganism in patients and healthy people including samples (*Acinetobacter sp P signifies that several species of Acinetobacter were present in the specimens). Microorganisms Patients Healthy p-value Samples Samples Total wounds Samples p-value Percentage people (Patients/Heal Wound edges Wound centers (edge / center) Healthy skins (Wounds / Healthy (Number of thy people) skins) positive/Number of tested) Pseudomonas aeruginosa 80% (16/20) 9% (1/11) ˂ 0.001 64.2% (27/42) 69 % (29/42) 66.6 % (56/84) 4.5% (1/22) ˂ 0.001 Staphylococcus aureus 60% (12/20) 0% (0/11) ˂ 0.001 40.4 % (17/42) 28.5 % (12/42) 34.5 % (29/84) 0% (0/22) ˂ 0.001 Streptococcus pyogenes 30% (6/20) 9% (1/11) 0.1 23.8 % (10/42) 11.9 % (5/42) 17.8% (15/84) 4.5% (1/22) 0.06 Acinetobacter spp. 80% (16/20) 0% (0/11) ˂ 0.001 61.9% (26//42) 50% (21//42) 55.9 % (47/84) 0% (0/22) ˂ 0.001 "Acinetobacter sp P" 55% (11/20) 0% (0/11) ˂ 0.001 35.7% (15/42) 28.5% (12/42) 32.1% (27/84) 0% (0/22) ˂ 0.001 Acinetobacter baumannii 10% (2/20) 0% (0/11) 0.2 7.1% (3/42) 9.5% (4/42) 8.3 % (7/84) 0% (0/22) 0.09 Acinetobacter nosocomialis 10% (2/20) 0% (0/11) 0.2 9.5% (4/42) 2.3% (1/42) 5.9 % (5/84) 0% (0/22) 0.1 Acinetobacter 5% (1/20) 0% (0/11) 0.3 4.7% (2/42) 4.7% (2/42) 4.7% (4/84) 0% (0/22) 0.1 guangdongensis 281 /285 Acinetobacter junii 5% (1/20) 0% (0/11) 0.3 4.7% (2/42) 0% (0/42) 2.3 % (2/84) 0% (0/22) 0.3 Acinetobacter lwoffii 5% (1/20) 0% (0/11) 0.3 0% (0/42) 4.7% (2/42) 2.3 % (2/84) 0% (0/22) 0.3 Rickettsia felis 5% (1/20) 0% (0/11) 0.3 0% (0/42) 2.3% (1/42) 1.9 % (1/84) 0% (0/22) 0.3 Acinetobacter junii and Acinetobacter lwoffii Mansonella spp., T. pallidum, and Poxvirus were also reported that wound chronicity is not were identified in the same patient (5%, 1/20 not detected in either wounds or healthy skin. associated with a single species of bacteria but versus 0%; p=0.6). There was 99% similarity with rather to a polymicrobial biofilm formed by the A. junii strain NBRC109759 (LC102684) and bacteria (Dowd et al., 2008; Percival et al., 100% similarity with the A. lwoffii strain DISCUSSION 2010). LMG1300 (EF611398). A. junii was detected in In this study, we evaluate the prevalence of two of the 42 specimens (4.7%) from the edges The skin is mainly colonized by non-pathogenic several microorganisms in skin samples taken of wounds. A. lwoffii was detected in two of the bacterial flora. In healthy people, some from people with chronic wounds and healthy 42 specimens (4.7%) from the center of wounds pathogenic bacteria, particularly S. aureus, can people, using molecular methods in Guinea (Figure 2). colonize the skin without clinical manifestations (Conakry), Africa. Our data are consistent as Identification failed for the 26 others samples (Wertheim et al., 2005). Asymptomatically, S. they are based on rigorous interpretation criteria. which were tested positive for Acinetobacter spp. aureus colonization is estimated to affect The quality of DNA extracts was systematically A mixed electropherogram indicating approximately 30% of the human population checked and each PCR assay was also polymicrobial infection with several Acinetobacter (Tong et al., 2015). In addition, S. aureus can systematically validated by the presence of species was observed for all these samples cause various diseases, including skin and soft positive and negative controls. In addition, each (Figure 3). Polymicrobial Acinetobacter infection tissue infections, particularly when skin or sample which tested positive for a was observed in 11 of the 20 patients (55%), mucosal barriers have been breached (Tong et microorganism with an initial PCR assay was marking a significant difference compared to al., 2015). Indeed, rupture of the skin barrier is systematically confirmed by a second PCR healthy people (0%, 0/11; p<0.001). the primary factor promoting infection (Scales assay targeting a sequence other than that S. pneumoniae, C. burnetii, Salmonella spp., and Huffnagle, 2013) allowing microorganisms to previously tested. Mycobacteria spp., H. ducreyi, Leishmania spp., enter, multiply, and spread within the body. It is Ehounoud et al. 033 Figure 2. Phylogenetic tree highlighting the position of Acinetobacter species identified in the study. The Rpo B sequences were aligned using MEGA 6 and phylogenetic inferences were obtained using the maximum likelihood standard method. A/ Example 1 B/ Example 2 Figure 3. Two examples of mixed electropherogram observed in the study. They indicate polymicrobial infection with several Acinetobacter species. 282 / 285 034. Glo. Adv. Res. J. Microbiol. As previously reported, S. aureus and P. aeruginosa case, it remains unknown whether R. felis from L. were the most common bacteria identified in chronic bostrychophila identified in wounds is merely a wounds, with a significant difference compared to healthy contamination by insect parts or whether, once inoculated, skin (Gjødsbøl et al., 2006; Rhoads et al., 2012; Serra et it plays a role in the infectious process in the wound. al., 2015). Each of these bacteria can express virulence Simultaneously, other pathogens which were tested, factors and surface proteins that may affect wound healing including Mycobacterium spp., S. pneumoniae, Salmonella (Serra et al., 2015; Bessa et al., 2015). Moreover, the co- spp., H. ducreyi, Leishmania spp., Mansonella spp., and infection of S. aureus and P. aeruginosa has also been poxvirus, were not detected. reported to be more virulent than a single infection. S. Overall, patients with chronic wounds present a mixed pyogenes was also highly prevalent in chronic wounds polymicrobial flora compared to the skin of healthy people. although there was no statistical significance to this. However, some results are not always as easy to explain. In our study, Acinetobacter spp., which has previously For example, Crisp et al. recently showed that the bacterial been reported in wounds, was significantly (80%) and cause of cellulitis cannot be determined by comparing the exclusively observed in patients with chronic wounds prevalence and quantity of pathogens from infected and (Gjødsbøl et al., 2006). Moreover, several species were uninfected skin biopsy specimens using current molecular identified, including for the same patient. Of the different techniques (qPCR and pyrosequencing) as well as species, A. baumannii was the most prominent causing standard culture techniques (Crisp et al., 2015). wide range of human infections (Rhoads et al., 2012). Indeed, A. baumannii is a well-known nosocomial pathogen with a high potential for antimicrobial resistance CONCLUSIONS (Eveillard et al., 2013). More recently, it has been involved in community-acquired infections and war- and natural The clinical management of chronic wounds is a real disaster-related infections, such as war wounds in troops challenge, particularly in patients with comorbidity and who from Iraq and Afghanistan (Eveillard et al., 2013). The live in poor areas. Our findings confirm that chronic implementation of molecular techniques has greatly wounds are colonized by multiple bacterial species as improved the identification of Acinetobacter species; this several bacterial species were observed in the skin, mainly may explain the diversity of Acinetobacter species from chronic wounds, in Guinea (Conakry). P. aeruginosa identified in wounds (A. baumannii, A. nosocomialis, A. and S. aureus were the more prevalent species identified. guangdongensis, A. junii, and A. lwoffii) (Rafei et al., 2014; Several different species of Acinetobacter were also Al Atrouni et al., 2016). In addition, two different species detected, including one, A. guangdongensis, which was were also identified, according to the area of the wound identified for the first time in skin. R. felis was also sampled (A. lwoffii from the edge and A. junii from the observed for the first time in this country. Chronic wounds center). All these species have been described as are colonized by many different bacteria which can originating from various environmental sources (Al Atrouni promote wound enlargement or delay healing. et al., 2016). They have previously been identified in human skin, with the exception of A. guangdongensis Declarations which was first described in 2014 from an abandoned lead- zinc ore mine (Feng et al., 2014; Al Atrouni et al., 2016). Ethics approval and consent to participate R. felis was also identified in the wound of one patient. This is the first time that this intracellular bacterium has This study was approved by the ethics committee of been identified in Guinea (Conakry). It had already been Guinea, Conakry (agreement number 008/CNERS/14). recovered from several countries in sub-Saharan Africa Written informed consent from all participants, including such as Senegal, Gabon, and Kenya/Tanzania, mainly patients and the parents or legal guardians of children was from the blood of febrile and afebrile patients (albeit more obtained. frequently in febrile patients). R. felis has also been reported in eschars in Senegal as well as in healthy skin Consent for publication swabs (Mediannikov et al., 2013; Mediannikov et al., 2014). The first transmission route reported for R. felis was All authors have approved the manuscript for submission. through cat fleas, Ctenocephalides felis. Recently, R. felis was detected in mosquitoes. It has been also Availability of data and materials demonstrated that Anopheles gambiae mosquitoes may be a vector of the bacterium (Dieme et al., 2015). Another All relevant data are within the paper. possibility is that the wound may be contaminated by the R. felis from environmental sources as booklice, Liposcelis bostrychophila, which is systematically infected by R. felis (Parola et al., 2015; Angelakis et al., 2016). If this is the 283 / 285 Ehounoud et al. 035 Competing interests Essayagh M, Essayagh T, Essayagh S, El Hamzaoui S (2014). Epidemiology of burn wound infection in Rabat, Morocco: Three-year review. Med Sante Trop.24:157-164. The authors declare that they have no competing interests. Eveillard M1, Kempf M, Belmonte O, Pailhoriès H, Joly-Guillou ML (2013). Reservoirs of Acinetobacter baumannii outside the hospital and Funding potential involvement in emerging human community-acquired infections. Int J Infect Dis.17:e802-5. Feng GD, Yang SZ, Wang YH, Deng MR, Zhu HH (2014). Acinetobacter This study was supported by Méditerranée Infection and guangdongensis sp. nov., isolated from abandoned lead-zinc ore. Int J the National Research Agency under the program Syst Evol Microbiol.64:3417-21. « Investissements d’avenir », reference ANR-10-IAHU-03. Ferdouse F, Hossain MA, Paul SK, Ahmed S, Mahmud MC, Ahmed R, Mahmud MdC, R. Ahmed, Haque AKMF, Nur-a-Alam Khan M, Ghosh S, Urushibara N, Kobayashi N (2015). Rickettsia felis Infection among Authors’ contributions Humans, Bangladesh, 2012-2013. Emerg Infect Dis. 21:1483-5. Gjødsbøl K, Christensen JJ, Karlsmark T, Jørgensen B, Klein BM, AKK, OM, and FF conceived and designed the framework Krogfelt KA (2006). Multiple bacterial species reside in chronic wounds: of this paper. CBE, AKK, AHB, AC, NA, JDN, and OM a longitudinal study, Int Wound J. 3:225-31. Gomez R, Murray CK, Hospenthal DR, Cancio LC, Renz EM, Holcomb performed the experiments. CBE, AKK, DR, OM, and FF JB, Wade CE, Wolf SE (2008). Causes of mortality by autopsy findings analyzed the data. CBE, AKK, AHB, AC, and JDN of combat casualties and civilian patients admitted to a burn unit. J Am contributed reagents, materials and analysis tools. CBE, Coll Surg. 208:348-54. DR, OM, and FF wrote the paper. All authors read and Gottrup F (2003). Organization of wound healing services: the Danish experience and the importance of surgery. Wound Repair Regen. 11: approved the final manuscript. 452-457. Guitard J, Edouard S, Lepidi H, Segonds C, Grare M, Ranty-Quintyn ML, Rouquette L, Cointault O, Rostaing L, Kamar N, Fenollar F (2012). ACKNOWLEDGEMENTS Identification of cause of posttransplant cachexia by PCR. Emerg Infect Dis.18:1386-8. Gulam-Abbas Z, Lutale JK, Morbach S, Archibald LK (2002). Clinical We are grateful to the infected and healthy volunteers who outcome of diabetes patients hospitalized with foot ulcers, Dar es agreed to participate in the study, as well as to Mamadou Salaam, Tanzania. Diabet Med. 19:575-579. Saliou Toure, Daouda Camara and the staff of Primary Hamad I, Forestier CL, Peeters M, Delaporte E, Raoult D, Bittar F (2015). Wild gorillas as a potential reservoir of Leishmania major. J Infect Health Care center of Maferinyah for technical assistance. Dis.211:267-73. Int Wound J. 12:47-52. Keita AK, Dubot-Pérès A, Phommasone K, Sibounheuang B, REFERENCES Vongsouvath M, Mayxay M, Raoult D, Newton PN, Fenollar F (2015). High prevalence of Tropheryma whipplei in Lao kindergarten children. Al Atrouni A, Joly-Guillou ML, Hamze M, Kempf M (2016). Reservoirs of PLoS Negl Trop Dis.9:e0003538. Non-Acinetobacter baumannii Species. Front Microbiol.7:49. La Scola B, Gundi VA, Khamis A, Raoult D (2006). Sequencing of the Angelakis E, Mediannikov O, Parola P, Raoult D (2016). Rickettsia felis: rpoB gene and Flanking spacers for molecular identification of The complex journey of an emergent human pathogen. Trends Acinetobacter species. J Clin Microbiol. 44:827-32. Parasitol. 32:554-64. Lavender CJ, Globan M, Johnson PD, Charles PG, Jenkin GA, Ghosh N, Benotmane A, Mohammedi F, Ayad F, Kadi K, Azzouz A (2000). Foot Clark BM, Martinello M, Fyfe JA (2012). Buruli ulcer disease in travelers lesions: etiologic and prognostic factors. Diabetes Metab. 26:113-117. and differentiation of Mycobacterium ulcerans strains from Northern Bessa LJ, Fazii P, Di Giulio M, Cellini L (2015). Bacterial isolates from Australia. J Clin Microbiol. 50:3717-21. infected wounds and their antibiotic susceptibility pattern: some Leslie DE, Azzato F, Karapanagiotidis T, Leydon J, Fyfe J (2007). remarks about wound infection. Development of a real-time PCR assay to detect Treponema pallidum Bouvresse S, Socolovshi C, Berdjane Z, Durand R, Izri A, Raoult D, in clinical specimens and assessment of the assay's performance by Chosidowa O, Brouqui P (2011). No evidence of Bartonella quintana comparison with serological testing. J Clin Microbiol. 45:93-6. but detection of Acinetobacter baumannii in head lice from elementary Mediannikov O, Fenollar F, Socolovschi C, Diatta G, Bassene H, Molez school children in Paris. Comp Immunol Microbiol Infect Dis.34:475-7. JF, Sokhna C, Trape JF, Raoult D (2010). Coxiella burnetii in humans Crisp JG, Takhar SS, Moran GJ, Krishnadasan A, Dowd SE, Finegold and ticks in rural Senegal. PLoS Negl Trop Dis.4:e654. SM, H. Summanen H, Talan DA (2015). Inability of polymerase chain Mediannikov O, Socolovschi C, Edouard S, Fenollar F, Mouffok N, reaction, pyrosequencing, and culture of infected and uninfected site Bassene H, Diatta G, Tall A, Niangaly H, Doumbo O, Lekana-Douki JB, skin biopsy specimens to identify the cause of cellulitis. Clin Infect Znazen A, Sarih M, Ratmanov P, Richet H, Ndiath MO, Sokhna C, Dis.61:1679-87. Parola P, Raoult D (2013). Common epidemiology of Rickettsia felis Dieme C, Bechah Y, Socolovschi C, Audoly G, Berenger JM, Faye O infection and malaria, Africa. Emerg Infect Dis. 19:1775-83. (2015). Transmission potential of Rickettsia felis infection by Anopheles Mediannikov O, Socolovschi C, Million M, Sokhna C, Bassene H, Diatta gambiae mosquitoes. Proc Natl Acad Sci U S A.112:8088-93. G, Fenollar F, Raoult D (2014). Molecular identification of pathogenic Dowd SE, Wolcott RD, Sun Y, McKeehan T, Smith E, Rhoads D (2008). bacteria in eschars from acute febrile patients, Senegal. Am J Trop Polymicrobial nature of chronic diabetic foot ulcer biofilm infections Med Hyg. 91:1015-1019. determined using bacterial tag encoded FLX amplicon pyrosequencing Mourembou G, Fenollar F, Lekana-Douki JB, Ndjoyi Mbiguino A, (bTEFAP). PLoS One.3:e3326. Maghendji Nzondo S, Matsiegui PB, Manego RZ, Ehounoud CH, Bittar Ehounoud CB, Yao KP, Dahmani M, Achi YL, Amanzougaghene N, F, Raoult D, Mediannikov O (2015b). Mansonella, including a Potential Kacou N'D, N’Guessan JD, Raoult D, Fenollar F, Mediannikov O New Species, as Common Parasites in Children in Gabon. PLoS Negl (2016). Multiple Pathogens Including Potential New Species in Tick Trop Dis.9:e0004155. Vectors in Côte d'Ivoire. PLoS Negl Trop Dis.10:e0004367. 284 / 285 036. Glo. Adv. Res. J. Microbiol. Mourembou G, Lekana-Douki JB, Mediannikov O, Nzondo SM, Kouna LC, Serra R, Grande R, Butrico L, Rossi A, Settimio UF, Caroleo B, Amato B, Essone JC, Biteghe Bi, Fenollar F, Raoult D (2015a). Possible Role of Gallelli L, de Franciscis S (2015). Chronic wound infections: the role of Rickettsia felis in Acute Febrile Illness among Children in Gabon. Pseudomonas aeruginosa and Staphylococcus aureus. Expert Rev Anti Emerg Infect Dis. 21:1808-15. Infect Ther.13:605-13. Mourembou G, Nzondo SM, Ndjoyi-Mbiguino A, Lekana-Douki JB, Kouna Smith A (2004). Etiology of the Problem Wound. In: Sheffield PJ, Fife CE, LC, Matsiegui PB, Manego RZ, Moukandja IP, Keïta AK, Tissot-Dupont Smith APS, eds. Wound Care Practice, Flagstaff: Best Publishing H, F. Fenollar F, Raoult D (2016). Cocirculation of Plasmodium and Company. 3-48. Bacterial Deoxyribonucleic Acids in Blood of Febrile and Afebrile Socolovschi C, Mediannikov O, Sokhna C, Tall A, Diatta G, H. Bassene H, Children from Urban and Rural Areas in Gabon, Am J Trop Med Hyg. Trape JF, Raoult D (2010). Rickettsia felis, a common cause of 95:123-32. uneruptive fever in rural Senegal. Emerg Infect Dis. 16:1140-2. Parola P, Mediannikov O, Dieme C, Raoult D (2015). Reply to Slesak, et Thomsen TR, Aasholm MS, Rudkjobing VB, Saunders AM, Bjarnsholt T, al: So much about Rickettsia felis infection to be discovered. Proc Natl Givskov M, Kirketerp-Møller MK, Nielsen PH (2010). The bacteriology Acad Sci U S A. 112:6595-6. of chronic venous leg ulcer examined by culture-independent molecular Percival SL, Thomas JG, Williams DW (2010). Biofilms and bacterial methods. Wound Repair Regen.18:38-49. imbalances in chronic wounds: anti-Koch. Int Wound J.7:169-75. Tong SY, Davis JS, Eichenberger E, Holland TL, Fowler VG Jr (2015). Pratt B, Cheesman J, Breslin C, Do MT (2016). Occupational injuries in Staphylococcus aureus infections: epidemiology, pathophysiology, Canadian youth: an analysis of 22 years of surveillance data collected clinical manifestations, and management. Clin Microbiol Rev.28:603- from the Canadian Hospitals Injury Reporting and Prevention Program. 61. Health Promot Chronic Dis Prev Can. 36:89-98. Wansbrough-Jones M, Phillips R (2006). Buruli ulcer: emerging from Rafei R, Kempf M, Eveillard M, Dabboussi F, Hamze M, Joly-Guillou ML obscurity. Lancet. 367:1849-58. (2014). Current molecular methods in epidemiological typing of Wertheim HF, Melles DC, Vos MC, van Leeuwen W, van Belkum A, Acinetobacter baumannii. Future Microbiol. 9:1179-94. Verbrugh HA, Nouwen JL (2005). The role of nasal carriage in Rhoads DD, Wolcott RD, Sun Y, Dowd SE (2012). Comparison of culture Staphylococcus aureus infections. Lancet Infect Dis. 5:751-62. and molecular Identification of bacteria in chronic wounds. Int J Mol Sci. Wolcott RD, Cox SB, Dowd SE (2010b). Healing and healing rates of 13: 2535-2550. chronic wounds in the age of molecular pathogen diagnostics. J Wound Rolain JM, Raoult D (2005). Molecular detection of Coxiella burnetii in Care. 19:272-8, 280-1. blood and sera during Q fever. QJM. 98:615-7; author reply 617-20. Wolcott RD, Rhoads DD, Bennett ME, Wolcott BM, Gogokhia L, Costerton Rolain JM, Sthul L, Maurin M, Raoult D (2002). Evaluation of antibiotic JW, Dowd SE (2010a). Chronic wounds and the medical biofilm susceptibilities of three rickettsial species including Rickettsia felis by a paradigm. J Wound Care. J Wound Care. 19:45-6, 48-50, 52-3. quantitative PCR DNA assay. Antimicrob Agents Chemother.46: 2747– 2751. Sano D, Tieno H, Drabo Y, Sanou A (1998). Management of the diabetic foot, apropos of 42 cases at the Ougadougou University Hospital Center. Dakar Med. 43:10-113. Scales BS, Huffnagle GB (2013).The microbiome in wound repair and tissue fibrosis. J Pathol. 229:323-31. Sen CK, Gordillo GM, Roy S, Kirsner R, Lambert L, Hunt TK, Gottrup F, Gurtner GC, Longaker MT (2009). Human skin wounds: a major and snowballing threat to public health and the economy. Wound Repair Regen. 17:763-71. 285 / 285