Phylogeny and Systematics of Panaspis and Afroablepharus Skinks

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2015-01-01 Phylogeny and Systematics of Panaspis and Afroablepharus Skinks (Squamata: Scincidae) in the Savannas of Sub-Saharan Africa Maria Fernanda Medina University of Texas at El Paso, [email protected]

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Recommended Citation Medina, Maria Fernanda, "Phylogeny and Systematics of Panaspis and Afroablepharus Skinks (Squamata: Scincidae) in the Savannas of Sub-Saharan Africa" (2015). Open Access Theses & Dissertations. 1099. https://digitalcommons.utep.edu/open_etd/1099

This is brought to you for free and open access by DigitalCommons@UTEP. It has been accepted for inclusion in Open Access Theses & Dissertations by an authorized administrator of DigitalCommons@UTEP. For more information, please contact [email protected]. PHYLOGENY AND SYSTEMATICS OF PANASPIS AND AFROABLEPHARUS SKINKS

(SQUAMATA: SCINCIDAE) IN THE SAVANNAS OF SUB-SAHARAN AFRICA

MARIA FERNANDA MEDINA

Department of Biological Sciences

APPROVED:

______Eli B. Greenbaum, Ph.D. Chair

______Carl Lieb, Ph.D.

______Chuan Xiao, Ph.D.

______Charles Ambler, Ph.D. Dean of the Graduate School Copyright © by Maria F. Medina

2015 PHYLOGENY AND SYSTEMATICS OF PANASPIS AND AFROABLEPHARUS SKINKS

(SQUAMATA: SCINCIDAE) IN THE SAVANNAS OF SUB-SAHARAN AFRICA

by

MARIA FERNANDA MEDINA, B.S.

THESIS

Presented to the Faculty of the Graduate School of

The University of Texas at El Paso

in Partial Fulfillment

of the Requirements

for the Degree of

MASTER OF SCIENCE

Department of Biological Sciences

THE UNIVERSITY OF TEXAS AT EL PASO

May 2015 ACKNOWLEDGMENTS

First and foremost, I would like to thank my parents for making sacrifices to give me a better future than they could’ve wished for. For encouraging me to reach beyond expectations and assuring me that family is my center, regardless of distance. As a young immigrant, it was very difficult to acclimate to a new place away from home. I also thank my significant other,

Nick Rodriguez, for giving me continual personal and academic support. If it weren’t for him, I wouldn’t have grown into the person I am now.

I want to express the deepest appreciation to my committee chair, Dr. Eli Greenbaum, for taking me under his wing even though I lacked basic knowledge of genetics. I am grateful for his incessant patience, passion for his work, and sincere interest in the growth of his students.

Thanks to his guidance, I am inspired to pursue a DVM/PhD degree in Veterinary Medicine. I thank my committee member and former Biochemistry professor. Dr. Xiao, for giving me a rigorous yet enthralling education on the subject. I also thank Dr. Lieb, for mentoring and patiently training me in skink biology and anatomy.

In addition, a thank you to my lab colleagues and true friends, Chris Anderson, Frank

Portillo, Daniel Hughes and Thornton Larson for support and advice with analyses and the research process, Nancy Conkey for teaching me lab procedures, and Waleeja Rashid and

Samantha Stewart for lab assistance and support.

iv ABSTRACT

Snake-eyed skinks are relatively small lizards of the genera Panaspis and Afroablepharus. Along with closely-related scincid taxa, the allocation of these genera has been constantly rearranged based on morphology, ecology and biogeography. Members of these genera occur primarily in savanna habitats throughout sub-Saharan Africa and include species that have highly conserved morphology, which poses a challenge for taxonomic studies. We sequenced two mitochondrial

(16S and cyt b) and two nuclear genes (PDC and RAG1) from 91 Panaspis and Afroablepharus samples from various localities in eastern, central and southern Africa. Concatenated gene-tree and divergence dating analyses were conducted to infer phylogenies and biogeographic patterns.

Molecular data sets revealed several cryptic lineages, with most radiations occurring during the mid-Miocene to Pliocene. We infer that rifting processes (including the formation of the East

African Rift System) and climatic oscillations contributed to the expansion of savannas and precipitated cladogenesis in snake-eyed skinks. Species in Panaspis and Afroablepharus used in this study formed a monophyletic group, rendering Afroablepharus as synonymous to Panaspis.

As a result, species previously assigned to Afroablepharus must be allocated to Panaspis or a different genus. Divergence estimates and cryptic speciation patterns of snake-eyed skinks were consistent with previous studies of other savanna vertebrate lineages from the same areas examined in this study.

v TABLE OF CONTENTS

ACKNOWLEDGMENTS ...... iv

ABSTRACT...... v

TABLE OF CONTENTS...... vi

LIST OF TABLES...... viii

LIST OF FIGURES ...... ix

CHAPTER 1: INTRODUCTION...... 1

1.1 Background ...... 1

1.2 Natural History...... 4

1.3 Taxonomy and Systematics...... 5

1.4 Research Objectives ...... 11

CHAPTER 2: MATERIALS AND METHODS ...... 12

2.1 Specimen acquisition...... 12

2.2 DNA extraction ...... 12

2.3 DNA amplification, sequencing and alignment ...... 14

2.4 Phylogenetic analyses...... 15

2.5 Divergence time estimation...... 16

CHAPTER 3: RESULTS...... 18

3.1 Phylogenetic analyses...... 18

3.2 Divergence time estimation...... 21

CHAPTER 4: DISCUSSION...... 26

4.1 Biogeography ...... 26

vi 4.2 Divergence dating...... 30

4.3 Taxonomy and species limits ...... 33

4.4 Conservation...... 37

LITERATURE CITED ...... 39

APPENDIX I ...... 53

VITA...... 60

vii LIST OF TABLES

Table 2.3.1 Primer sequences used in this study...... 15

viii LIST OF FIGURES

Figure 1.3.1. Map of central, eastern and southern Africa showing the disjunct distribution of

Panaspis wahlbergi (in orange). Sampled localities for the genetic analyses are shown with circles. Circle colors correspond to the clades in Fig. 3.1.1. Map was modified from Branch

(1998) and Spawls et al. (2002). Black circles indicate type localities of Ablepharus anselli

(Kasempa, Zambia), Ablepharus moeruensis (Kilwa Island, Lake Mweru between Zambia and

Katanga, DRC) and Panaspis seydeli (Lubumbashi, southeast Katanga, DRC). All of these taxa are currently considered to be synonyms of Afroablepharus seydeli (Uetz and Ho!ek, 2015).

...... 10

Figure 3.1.1. Maximum likelihood tree derived from 16S, cyt b, PDC and RAG1 DNA

sequences. Nodes supported by Bayesian posterior probability of " 0.95 and maximum

likelihood bootstrap support of " 70 are indicated by black circles. Nodes supported by

maximum likelihood values of " 70 only are indicated by open circles. Picture (MBUR 7835)

depicts skink in the A. wahlbergi complex...... 20

Figure 3.2.1. Mitochondrial (cyt b) phylogenetic tree from BEAST analyses. Open circles denote

calibration points, and numbers at the base of nodes denote mean highest posterior densities

(HPD)...... 22

Figure 3.2.2. Nuclear (RAG1) phylogenetic tree from BEAST analyses. Open circles denote

calibration points, and numbers at the base of nodes denote mean highest posterior densities

(HPD)...... 24

ix CHAPTER 1: INTRODUCTION

1.1 Background

There are about 9671 living species of squamates accounting for a great part of the world’s biodiversity. The major groups (Anguimorpha, Gekkota, Iguania, Lacertibaenia,

Serpentes, Scinciformata and Teiformata) of squamates likely diversified during the Jurassic

Period about 200–150 million years ago (mya). Further radiations during the Cenozoic (50 mya) generated the different squamate families that are recognized today (Vidal and Hedges, 2009).

Formerly part of the group Scleroglossa, skinks were distinguished from lizards of the group

Iguania by prehension of prey with their teeth and jaws (Iguanians use their muscular tongues).

This scleroglossan adaptation expanded the tongue’s chemosensory capabilities, which allowed the group to modify eating patterns in response to their prey and exploit new adaptive zones around the world (Schwenk, 2000). The Scleroglossa are no longer supported. Vidal and Hedges

(2009), using a phylogeny constructed with nuclear protein-coding genes, separated a new grouping within Squamata that found jaw prehension to be an ancestral trait. Scinciformata is thus a cosmopolitan lineage (excluding Antarctica) composed of the families Cordylidae,

Gerrhosauridae, Scincidae, and Xantusiidae, all of which are now grouped under Unidentata (one egg tooth). Iguanians are now positioned as the sister group to a clade including snakes and anguimorphs.

The Family Scincidae is thought to have radiated very early in the Cenozoic era, around the time of a major asteroid impact at the end of the Cretaceous ca. 65 mya. This event precipitated ecological changes and various extinctions at a global level (Vidal and Hedges,

2009). According to fossil records from Australia and South Africa, lizards originated in

Gondwana, the southern land mass that split off from Pangaea 135 mya (Pianka and Vitt, 2003).

1 Historically, the use of skull morphology separated the Family Scincidae into four subfamilies:

Acontinae, Feylininae, Lygosominae and Scincinae (Greer, 1970). Although Panaspis and related genera were considered lygosomine, recent molecular and morphological evidence

(Hedges, 2014; Hedges and Conn, 2012; Skinner et al., 2011) supported the division of skinks into nine families, of which seven (Acontinae, Egerniinae, Eugongylinae, Lygosominae,

Mabuyinae, Sphenomorphinae and Scincinae) are currently recognized as subfamilies (Uetz and

Ho!ek, 2015). By this classification, the genera Afroablepharus, Panaspis, Lacertaspis and

Leptosiaphos are currently allocated to the subfamily Eugongylinae (Uetz and Ho!ek, 2015).

According to Uetz and Ho!ek (2015), the African scincid genus Panaspis includes 8 species: P. breviceps (Peters, 1873), P. burgeoni (de Witte, 1933b), P. cabindae (Bocage, 1866),

P. helleri (Loveridge, 1932), P. megalurus (Nieden, 1913), P. nimbaensis (Angel, 1944), P. tancredi (Boulenger, 1909) and P. togoensis (Werner, 1902). The genus Afroablepharus, formerly a subspecies to Panaspis, contains 7 species: A. africanus (Gray, 1845), A. annobonensis (Fuhn, 1972), A. duruarum (Monard 1949), A. maculicollis Jacobsen and

Broadley, 2000, A. seydeli (de Witte, 1933a), A. wahlbergi (Smith, 1849) and A. wilsoni

(Werner, 1914).

Panaspis was created by Cope (1868) specifically for his new species anaeus, which was later synonymized with Ablepharus cabindae (Bocage, 1866). As a result, Panaspis was synonymized with Ablepharus Lichtenstein 1823 by Boulenger (1887), and later with Riopa

Gray, 1839 by Smith (1937). The latter author was ignored despite drawing attention to similarities in head scalation and distinctive characters in the lower eyelid. Smith’s methodology and Mittleman’s (1952) strong disbelief in subgenera led him to revise all the genera in the subfamily Lygosominae, resulting in the validation of Panaspis as a full genus. Greer and Parker

2 (1967) introduced the use of skull characteristics when they described a scincid lizard from the island of Bougainville (Solomon Islands). Three years later, Greer would apply skull structure to arrange the scincid subfamilies Acontinae, Feylininae, Lygosominae and Scincinae (Greer,

1970).

As mentioned earlier, recent molecular and morphological evidence (Hedges, 2014;

Hedges and Conn, 2012; Skinner et al., 2011) has suggested the division of skinks into nine families. Although considered controversial (Pyron et al., 2013), this new subdivision supports the monophyly of Scincidae as a currently accepted preliminary hypothesis (Uetz and Ho!ek,

2015). The genus Panaspis has been rearranged several times, focusing on eyelid structure, morphology of the skull pterygoids and palates, and cephalic lepidosis (Broadley, 1989; Fuhn,

1969, 1972; Greer, 1974; Perret, 1973, 1975, 1982). As a result, different species were moved back and forth between different genera, including Leptosiaphos and Afroablepharus (Greer,

1974; Perret, 1973, 1975). Currently, Panaspis includes all African species with lower-mobile eyelids or pre-ablepharine eyes (lower eyelid fused or with a palpebral slit, respectively). African skink species with ablepharine eyes are allocated to the genus Afroablepharus.

This study started as an undergraduate research project, which focused on comparing specimens from Katanga Province, Democratic Republic of the Congo (DRC) with the widespread species Afroablepharus wahlbergi. A preliminary phylogeny based on DNA sequence data confirmed that the Katanga specimens, as well as borrowed specimens from other museums and herpetologists, were not congruent with A. wahlbergi. Instead, cryptic speciation of at least 20 clades was revealed. Therefore, this study focused on identifying all obtained specimen and revising the relationships between Panaspis and Afroablepharus.

3 1.2 Natural History

Panaspis, commonly known as snake-eyed skinks for their inability to blink, are relatively small lizards (adult size less than 70 mm total length) that inhabit lowland savannas, woodlands and forests in sub-Saharan Africa. Most species occur in savannas and woodlands; only P. breviceps inhabits lowland rainforests at elevations up to 800 meters (Chirio and

LeBreton, 2007). Panaspis and Afroablepharus skinks are small, diurnal, pentadactyle, slender, and their tail measures a little more than half their total length (Pianka and Vitt, 2003; Largen and Spawls, 2010). Their smooth scales are usually arranged in 22–28 rows around the middle of the body (Largen and Spawls, 2010).

Panaspis skinks were known for having an “ablepharine” eye—except for P. cabindae, with a pre-ablepharine eye—with fused eyelids and a transparent disk as in the eye of a snake.

However, all ablepharine skinks are currently allocated to the genus Afroablepharus, leaving P. cabindae and skinks that have lower-mobile eyelids in the genus Panaspis (Uetz and Ho!ek,

2015). Mobile eyelids in Panaspis and closely related skinks (e.g., Leptosiaphos) tend to have a transparent “window” that allows them to see when their eyes are closed. Panaspis and

Afroablepharus are mostly semifossorial, dwelling mainly under leaf litter, but also found in termitaria, grassy spots, or under rocks and rotting logs (FitzSimons, 1943). In describing P. cabindae, Schmidt (1919; 566) stated, “only on searching through and separating the grass can these skinks be observed moving about swiftly, more in the fashion of snakes than lizards.”

Because of their semifossorial nature, they feed on burrowing insects such as termites (Schmidt,

1919). Afroablepharus wahlbergi is known to feed on spiders, beetles and their larvae, cockroaches, caterpillars, woodlice and ants (FitzSimons, 1943; Razzetti, 2002) when termites are not available. Spawls et al. (2002) stated that males in South Africa live 10 to 12 months and

4 the females live a few years longer. Sexual maturity is reached at 8 to 9 months, and males develop a pinkish-orange pigmentation on the chin during the breeding season (Largen and

Spawls, 2010). About 2–6 eggs are laid between November and January under stones or logs after mating takes place in the late summer (Branch, 1998). Hatchling size ranges from 30–35 mm total length, and the eggs measure roughly 4 mm x 7 mm (FitzSimons, 1943; Spawls et al.,

2002). The coloration pattern of Panaspis and Afroablepharus skinks consists of a pale gray, with several dark lines along the brown or bronze dorsum and/or flanks, “pale dorsolateral stripes”, and a dark brown or black lateral band that sometimes exhibits “light mottling”, either scattered, arranged in a dotted line or diagonal rows of three dots (Jacobsen and Broadley, 2010;

Largen and Spawls, 2010).

1.3 Taxonomy and Systematics

The complicated history of Panaspis is attributed to the repeated transfer of species between genera for many decades, and to the scarcity of morphological differences among putative species, especially when DNA analyses were not possible. The type species of the genera that are most involved in the various allocations of snake-eyed African skinks into

Panaspis are the following: Ablepharus (A. pannonicus Fitzinger, 1924), Afroablepharus

(Cryptoblepharus wahlbergi Smith, 1849), Lacertaspis (L. reichenowii Peters, 1874),

Leptosiaphos (L. meleagris Boulenger, 1907) and Panaspis (P. cabindae Cope, 1868).

Ablepharus is the oldest genus of the group and is now restricted to skinks from southeastern

Asia (Uetz and Ho!ek, 2015). The remaining genera were created during the subsequent revisions of Panaspis skinks by later herpetologists.

5 Cope (1868) erected the genus Panaspis after naming his new species anaeus in Angola.

Similarities in eye structure—no movable eyelids, a transparent disc covering the eye—led

Boulenger (1887) to synonymize Panaspis with Ablepharus. Fuhn (1969) revised this action and suggested that Panaspis and other genera be separated from Ablepharus, as he believed that the trait of the ablepharine eye was not thoroughly examined. He found some ablepharine skinks with a palpebral slit hidden under their supercilium [arch above the eye], which suggested that not all Ablepharus had fused, unmovable eyelids and a transparent disk; in fact, a movable lower eyelid with a transparent scale was present. Because Fuhn found that the osteological pattern between the lizards’ skulls was more convincing than eye anatomy, he assigned Ablepharus to

Eurasian species only, and restricted Panaspis to skinks from Africa (cabindae, duruarum, megalurus, seydeli, tancredi, wahlbergi and wilsoni). Consequently, he suggested further studies on skull structure after Greer and Parker (1967) introduced the use of skull morphology to describe a new scincid species. Greer (1970) later conducted an extensive study that organized skink species in subfamilies (Acontinae, Feylininae, Lygosominae and Scincinae) based on their skull morphology, as well as ecological and geographical factors. Two years later, Fuhn (1972) included seven species that had a mobile lower eyelid with a transparent disc into Panaspis (P. africanus, P. breviceps, P. gemmiventris, P. kitsoni, P. nimbaensis, P. pauliani, P. reichenowi, P. rohdei, and P. vigintiserierum).

Perret (1973) added two more species he described (P. amieti and P. fuhni) to Panaspis, following the new identification key from Greer (1970). In the same study, Perret (1973) divided

Panaspis skinks into three groups based on morphology to improve his diagnosis of African scincids. These groups were: (1) Mabuiform, with an increased number of midbody scales and supranasals present; (2) Lacertiform, with a slender body, long extremities, an increased number

6 of subdigital lamellae under the fingers and toes, extremities overlapping when adpressed to the body, long and thin tail measuring more than twice the SVL, not enlarged at base; and (3)

Sepsinoïde, with a prolonged body, short and robust extremities that don’t overlap when adpressed to body and a muscular tail enlarged at base. According to his classification, the sepsinoïde group included the species amieti, fuhni, gemmiventris, lepesmei, pauliani, vigintiserierum, the lacertiform group included the species reichenowii and rohdei, and the mabuiform group contained africana, breviceps, kitsoni and nimbaensis. With this arrangement, only twelve species of Panaspis were recognized as full species.

To improve the most recent arrangement, Greer (1974) erected a new genus,

Afroablepharus, to accommodate species with an ablepharine eye (lower eyelid fixed and fused to the upper eyelid), and moved all species with movable lower eyelids and pre-ablepharine eyes—the latter referring to P. cabindae only—to Panaspis. He synonymized Leptosiaphos with

Panaspis (Greer, 1974) to satisfy the addition of the movable lower eyelid character. A year later, Perret (1975) erected the genus Lacertaspis to accommodate his lacertiform species, reichenowi and rohdei, but he decided to classify it as a subgenus along with Greer’s (1974)

Afroablepharus. Broadley (1989) thoroughly revised the genera in question and restricted

Panaspis to species residing in African savannas and having ablepharine or pre-ablepharine eyes.

He then restored Leptosiaphos to full genus rank for forest and montane grassland species that had a movable lower eyelid. He also erected a new subgenus, Perretia, for his new species P. rhomboidalis, which had a distinctive cephalic lepidosis. Recent revisions by Schmitz et al.

(2005) elevated Afroablepharus to full genus rank along with Lacertaspis and Leptosiaphos. The phylogeny of the latter authors demonstrated that P. wahlbergi belongs in the genus

Afroablepharus.

7 My current study on African and Asian skinks began after specimens were collected in

Katanga Province, DRC in a 2010 expedition by Eli Greenbaum (University of Texas at El

Paso). These specimens had morphology congruent with A. wahlbergi, one of the species known to reside in Katanga. The remaining four species that have been reported from Katanga,

Ablepharus anselli, A. moeruensis, P. seydeli and P. smithii, had different cephalic lepidosis.

After collecting molecular data from additional specimens from sub-Saharan Africa, I constructed a phylogeny that arranged all these samples in numerous distinct clades. Based on the relatively long branch lengths of lineages in my well-supported phylogeny, A. wahlbergi is a complex of at least 20 cryptic species. My analyses also confirmed that Panaspis forms a monophyletic clade with Afroablepharus, meaning the latter genus must be synonymized.

Divergence time estimations confirmed that most radiations of Panaspis/Afroablepharus occurred during the Miocene and Pliocene eras, when major geological and climatic changes across sub-Saharan Africa took place.

The status of P. seydeli and P. smithii is currently unclear. Panaspis seydeli is known from four specimens: one from Mwenzo in northeastern Zambia, and three type specimens that are currently synonymized with this species. These specimens include the type of Ablepharus anselli (from Kasempa, northwestern Zambia), the type of A. moeruensis (from Kilwa Island in

Lake Mweru, northeastern Zambia and southeastern DRC), and the type of P. seydeli (from

Lubumbashi, in southeastern Katanga, DRC) (Broadley and Cotterill, 2004; de Witte, 1933a).

Fuhn (1964, 1970) placed A. anselli in the synonymy of P. tancredii from Ethiopia, given the strong resemblance between the types of the two species. However, he suggested further revision because the two specimens implied an allopatric distribution; the populations are 3200 km apart, and the difference in elevation between both areas is approximately 1500 m (Broadley and

8 Cotterill, 2004; Largen and Spawls, 2006) (Fig. 1.3.1). Eventually, A. anselli, A. moeruensis and

P. tancredii were considered synonyms of P. seydeli (Jacobsen and Broadley, 2000) because of only slight differences in head scalation and midbody scale rows. However, Broadley and

Cotterill (2004) rejected this decision because of the evident biogeographic gap between P. tancredii and the other three species. Supporting this idea, Largen and Spawls (2006) resurrected

P. tancredii from the synonymy of P. seydeli, and provided a key that justified the separation.

Another concern is that P. smithii is also considered a synonym of P. seydeli, which is currently listed as Afroablepharus seydeli (Greer, 1974 in The Reptile Database (Uetz and Ho!ek, 2015).

However, P. smithii is still regarded as a distinct species by recent authors (Branch, 2008;

Broadley and Cotterill, 2004). Seeing that A. moeruensis was described in the Katanga area relatively far away from topotypic P. seydeli, it is possible that the two specimens belong to different taxa.

9

Figure 1.3.1. Map of central, eastern and southern Africa showing the disjunct distribution of Panaspis wahlbergi (in orange). Sampled localities for the genetic analyses are shown with circles. Circle colors correspond to the clades in Fig. 3.1.1. Map was modified from Branch (1998) and Spawls et al. (2002). Black circles indicate type localities of Ablepharus anselli (Kasempa, Zambia), Ablepharus moeruensis (Kilwa Island, Lake Mweru between Zambia and Katanga, DRC) and Panaspis seydeli (Lubumbashi, southeast Katanga, DRC). All of these taxa are currently considered to be synonyms of Afroablepharus seydeli (Uetz and Ho!ek, 2015).

10

1.4 Research Objectives

1. Identify topotypic Afroablepharus wahlbergi

2. Determine the extent of genetic diversity of Panaspis in sub-Saharan Africa

3. Identify the lineages revealed by my preliminary phylogeny

4. Estimate the biogeographic patterns that encouraged cryptic speciation in the A.

wahlbergi complex

5. Confirm the placement of Afroablepharus relative to Panaspis

6. Provide a species description for clades that represent new species

11

CHAPTER 2: MATERIALS AND METHODS

2.1 Specimen acquisition

The skink specimens that helped start this study were collected by Dr. Eli Greenbaum from Katanga Province, DRC during the wet season in January 2010. This study includes molecular and morphological data from the 2010 specimens and borrowed Afroablepharus cf. wahlbergi specimens from multiple localities in sub-Saharan Africa. Two species of Trachylepis

(Scincidae) and Cordylus marunguensis were used to root the trees. Additional sequences of closely related genera (Cordylus, Lacertaspis, Leptosiaphos, Mochlus and Xantusia) were also sequenced or included from GenBank.

2.2 DNA extraction

The DNA of tissue samples was extracted using the Qiagen DNeasy Blood and Tissue

Kit (Valencia, CA), and the IBI DNA Extraction Kit (Shelton Scientific, Peosta, IA). Small pieces of tissue of about 25 mg were cut, soaked in deionized water, and placed in a refrigerator at 2°C for two hours (h), allowing the ethanol to diffuse from the tissue. The samples were allowed to dry at 2ºC for 2 h. Once the 2 h drying period was completed, the process of the extraction kit of choice was initiated.

When using the Qiagen DNeasy Blood and Tissue Kit, 180 µl of Buffer ATL and 20 µl of

Proteinase K were added to the samples. Samples were then placed on a hot plate for 30 minutes

(min) at 55°C. After the 30 min incubation period, samples were removed, vortexed for 20 seconds (s), and then left overnight on the hot plate at 55°C. The next morning, 200 µl of Buffer

AL was added to each sample. Samples were vortexted for 20 s and placed on the hot plate for

10 min at 70°C. Following the 10 min incubation, 200 µl of 100% ethanol was added and each

12 sample was vortexed for 20 s. The resulting solution was placed into spin columns and centrifuged at 8000 revolutions per min (rpm) for 1 min. Liquid remaining in the spin columns was discarded and 500 µl of Buffer AW1 was added. Samples were again centrifuged at 8000 rpm for 1 min. Liquid remaining in the spin columns was discarded and 500 µl of Buffer AW2 was added. Samples were centrifuged at 14,000 rpm for 3 min. Liquid was again discarded, 200

µl of Buffer AE was added and allowed to incubate at room temperature for one min, and samples were then centrifuged at 8000 rpm for 1 min. The remaining solution was saved for polymerase chain reaction (PCR). The previous step was repeated to obtain a total of 400 µl of genomic DNA for PCR.

When using the IBI DNA Extraction Kit, a Micropestle was used to grind the tissue to a pulp. Then, 200 µl of GT Buffer were added. After continuing to grind each sample for a few seconds, 20 µl of Proteinase K were added to the sample mixture and vortexed for 20 s. The resulting mixtures were incubated at 60°C for 30 min to lyse the tissue, while inverting the tubes every 5 min. After incubation, 200 µl of GBT Buffer were added, and the samples were vortexed for 5 s. The samples were incubated at 60°C for 20 min until the sample lysate was clear. During incubation, the tubes were inverted every 5 min. After incubation, 200 µl of absolute ethanol were added to the clear lysate and vortexed for 10 s. Each sample mixture was transferred into a spin column and 2 mL collection tube. The spin columns were centrifuged at 14,000 rpm for 2 min. The liquid in the collection tube was discarded, and 400 µl of W1 Buffer was added to the spin column. The samples were then centrifuged at 14,000 rpm for 30 s, and the resulting liquid was again discarded. Then, 600 µl of Wash Buffer were added to the spin columns, and they were centrifuged at 14,000 rpm for 30 s. After discarding the liquid in the collection tubes, the samples were centrifuged again at 14,000 rpm for 3 min to dry the filter matrix in the spin

13 columns. The dried spin columns were transferred to clean 1.5 mL microcentrifuge tubes, and

200 µl of pre-heated Elution Buffer were added to each spin column. The samples were incubated at room temperature for 5 min, and then centrifuged at 14,000 rpm for 30 s. The remaining solution was saved for PCR. The previous step was repeated to obtain 200 µl of genomic DNA for PCR.

2.3 DNA amplification, sequencing and alignment

Two mitochondrial (16S and cyt b) and two nuclear (PDC and RAG1) genes were amplified in this study (primer sequences shown in Table 2.3.1). A negative control was used in all amplifications to ensure there is no contamination. Amplifications were completed with a denaturing temperature of 95°C for 2 minutes, followed by denaturation at 95°C for 35 seconds

(s), annealing at 50°C for 35 s, and extension at 72°C for 95 s with 4 s added to the extension per cycle for 32 or 34 cycles (for mitochondrial or nuclear genes, respectively). Results from PCR were visualized with 1.5% agarose gel and SYBRsafe gel stain (Invitrogen, Carlsbad, CA). The

PCR products were purified with Agencourt AMPure XP magnetic bead solution (Beckman

Coulter, Danvers, MA) following the manufacturer’s protocols. Forward and reverse strands of

PCR products were sequenced on an ABI 3700xl capillary DNA sequencer at the UTEP DNA

Core Facility. The program SeqMan (Swindell and Plasterer, 1997) was used to interpret chromatograph data. Sequences were aligned using the ClustalW algorithm in the program

MEGALIGN (Clewley and Arnold, 1997) and adjusted by eye in MacClade v4.08 (Maddison and Maddison, 2000). The latter program was also used to check the accuracy of protein-coding genes by translation to amino acids (Maddison and Maddison, 2000). Any hypervariable regions in 16S were removed from the final analysis.

14

Table 2.3.1. Primer sequences used in the study.

Primer Gene Reference Sequence 16L9 16S Pramuk et al. (2008) 5#-CGCCTGTTTACCAAAAACAT-3# 16H13 16S Pramuk et al. (2008) 5#-CCGGTCTGAACTCAGATCACGTA-3# CytbCBJ10933 cyt b Vences et al. (2003) 5'-TATGTTCTACCATGAGGACAAATATC-3' Cytbc cyt b Vences et al. (2003) 5'-CTACTGGTTGTCCTCCGATTCATGT-3' PHOF2 PDC Bauer et al. (2007) 5'-AGATGAGCATGCAGGAGTATGA-3' PHOR1 PDC Bauer et al. (2007) 5'-TCCACATCCACAGCAAAAAACTCCT-3' RAG1 G396 RAG1 Groth and 5’-TCTGAATGGAAATTCAAGCTGTT-3’ Barrowclough (1999) RAG1 G397 RAG1 Groth and 5’-GATGCTGCCTCGGTCGGCCACCTTT-3’ Barrowclough (1999) RAG1f700 RAG1 Bauer et al. (2007) 5'-GGAGACATGGACACAATCCATCCTAC-3' RAG1r700 RAG1 Bauer et al. (2007) 5'-TTTGTACTGAGATGGATCTTTTTGCA-3'

2.4 Phylogenetic analyses

The analysis included 91 individuals from the genera Panaspis and Afroablepharus, with various representatives of P. breviceps, P. togoensis, Afroablepharus africanus, A. annobonensis, P. cabindae, P. cf. tancredi, P. sp. Limpopo, P. sp. Namibia, A. maculicollis, P. sp. Mozambique 1, P. sp. Mozambique 2, P. sp. Malawi, P. sp. Mozambique 3, P. sp.

Mozambique 4, P. sp. Mozambique 5, P. sp. Gauteng, A. wahlbergi, P. sp. Mozambique 6, P. sp.

Tanzania 1, P. sp. Tanzania 2, P. sp. Katanga 1 and P. sp. Katanga 2. Outgroup samples included

Cordylus marunguensis, Trachylepis megalura, Trachylepis striata, Mochlus afer, Lacertaspis chriswildi, Lacertaspis reichenowi, Lacertaspis rhodei, Leptosiaphos blochmanni, Leptosiaphos gemmiventris, Leptosiaphos koutoui, Leptosiaphos meleagris and Leptosiaphos sp. We conducted phylogenetic analyses of single-gene and concatenated data sets, consisting of 2278 characters from the mitochondrial genes 16S (568 bp) and cyt b (619 bp) and nuclear genes PDC

(442 bp) and RAG1 (699 bp). Hypervariable regions in the 16S ribosomal gene, totaling 50 base

15 pairs, were removed from the final analysis. The program SeqMan (Swindell and Plasterer,

1997) was used to interpret chromatograph data. Sequences were aligned using the ClustalW algorithm in the program MEGALIGN (DNASTAR, Madison, WI, USA) and adjusted by eye in

MacClade v4.08 (Maddison and Maddison, 2000). The latter program was also used to check the accuracy of protein-coding genes by translation to amino acids. Sequence data were analyzed with maximum likelihood (ML) with the GTRGAMMA model in RAxML v7.2.6 (Stamatakis,

2006). All parameters were estimated, and a random starting tree was used. Node support was assessed with 1,000 nonparametric bootstrap replicates (Stamatakis et al., 2008). Bayesian inference (BI) was conducted with MrBayes 3.1 (Huelsenbeck and Ronquist, 2001; Ronquist and

Huelsenbeck, 2003). Our model included ten data partitions: a single one for 16S and independent partitions for each codon position of the protein-coding genes cyt b, PDC and

RAG1. Concatenated data sets were partitioned identically for ML and BI analyses. The Akaike information criterion in jModelTest 2 (Darriba et al., 2012) was used to find the model of evolution that was most consistent with our data for subsequent BI analyses. Bayesian analyses were conducted with random starting trees, run for 20,000,000 generations, and Markov chains were sampled every 1000 generations. Are we there yet? (AWTY) (Nylander et al., 2008) was used to verify that multiple runs converged. Once convergence was met, 25% of the trees were discarded as burn-in. Phylogenies were visualized using FigTree v1.4.2 (Rambaut, 2012).

2.5 Divergence time estimation

Because of the lack of calibration points for the age of Panaspis, Afroablepharus and related genera, we used secondary calibrations of taxa within the family Scincidae from Mulcahy et al. (2012). Sequences from squamate groups used by Mulcahy et al. (2012) were obtained

16 from GenBank (Appendix I). The program BEAST 1.8.0 (Drummond et al., 2012) was used on separate mitochondrial (cyt b, 619 bp) and nuclear (RAG1, 477 bp) data sets to estimate divergence dates of lineages within Panaspis and Afroablepharus. These data sets were run separately because of the lack of available matching sequences between outgroups, and to assess congruency in time estimations. The outgroup sequences we obtained from GenBank ranged from 379–1053 bp. Because other studies analyzed different regions of RAG1, we were only able to work with a relatively short overlapping fragment.

Secondary calibration points were assigned to extant clades and treated as minimum age constraints. We used the most recent common ancestor (MRCA) of Trachylepis/Tiliqua (33 million years [my]) and Lygosominae sensu lato (65 my) (Mulcahy et al., 2012). Ingroups that were assigned these calibration points were constrained as monophyletic. Outgroup sampling included cordylomorphs (Karusasaurus, Namazonurus, Smaug and Xantusia) and was also constrained. We used a lognormal relaxed-clock model with a Yule speciation tree prior. The mean and standard deviation were set to 1.0, and calibration points were entered as offsets.

Analyses were run for 50 million generations, sampling every 1,000 generations and discarding the first ten percent of generations as burn-in. Convergence and effective sample size for parameters were checked with Tracer v1.5 (Rambaut and Drummond, 2007). Mean ages and credibility intervals (CI) were calculated using TreeAnnotator 1.8 (Drummond et al., 2012). We visualized trees in FigTree 1.4.2 (Rambaut, 2012).

17

CHAPTER 3: RESULTS

3.1 Phylogenetic analyses

Our data set comprised 2278 base pairs (bp) from the mitochondrial genes 16S (518 bp) and cyt b (619 bp) and nuclear genes PDC (442 bp) and RAG1 (699 bp). Because of tissue degradation, 11 samples failed to amplify for the cyt b gene, 15 for the PDC gene, and 41 for the

RAG1 gene (Table 2.3.1). Other studies have shown that phylogenetic analyses with missing data can still be accurately inferred if they have an appropriate amount of informative characters.

Support actually improves when taxa with missing data are included, as opposed to excluding these taxa altogether (Jiang et al., 2014; Mulcahy et al., 2012; Pyron et al., 2013; Wiens and

Morrill, 2011). For the BI analyses, the models of nucleotide substitution selected by jModelTest

2 were the following: 16S (TIM2 + I + G); cyt b 1st codon position (TPM3uf + I + G), cyt b 2nd codon position (TPM2uf + G), cyt b 3rd codon position (TIM2 + I + G); PDC 1st codon position

(TIM3 + I), PDC 2nd codon position (TPM3uf + I), PDC 3rd codon position (TPM1uf + G);

RAG1 1st codon position (TPM1uf + G), RAG1 2nd codon position (HKY + G), RAG1 3rd codon position (TPM1 + G). The least restrictive model (GTR) was implemented whenever a model selected by jModelTest 2 was not available. The topologies for the ML and BI analyses were identical, and strong support values were similar for most clades (Fig. 3.1.1). These concatenated

ML and BI analyses resulted in the same topologies as our single-gene mtDNA analyses (not shown). The ML analysis likelihood score was –18445.331817.

The ML and BI analyses (Fig. 3.1.1) recovered strong support (> 70% ML bootstrap values and > 0.95 BI posterior probabilities) for a monophyletic clade including all Panaspis and

Afroablepharus samples, which is sister to a well-supported clade including Lacertaspis and

Leptosiaphos. Relationships among most clades within the former genera were also well

18 supported, and many of the 17 cryptic lineages identified from Katanga (DRC), Tanzania,

Angola, Malawi, Mozambique, Namibia, and South Africa are likely to be new species. Two monophyletic clades corresponding to P. togoensis and P. breviceps (known from forests and forest/savanna mosaic habitats) were recovered in a basal position to the remaining samples from savanna habitats, which form a well-supported clade. Within the latter group, a monophyletic clade including A. africanus, A. annobonensis and P. cabindae was recovered as basal to other samples from central, eastern and southern Africa. Based on a relatively long branch length, our only sample from western Ethiopia is a distinct lineage that corresponds to P. cf. tancredi. Other well-supported clades included: (1) P. sp. Limpopo in northern South Africa, (2) P. sp. Namibia,

(3) A. maculicollis from northern South Africa, (4) P. sp. Mozambique 1, in Gorongossa

National Park and provinces at the northwestern side of the country, (5) P. sp. Mozambique 2, located near the northeastern coast of the country, (6) P. sp. Malawi, (7) P. sp. Mozambique 3 in

Nampula Province, in northeastern Mozambique about 170 km south from the following lineage,

(8) P. sp. Mozambique 4 from Cabo Delgado Province in the northeastern side of the country,

(9) P. sp. Mozambique 5, located in east-central Sofala Province, (10) P. sp. Gauteng, from

Gauteng Province in east-central South Africa, (11) A. wahlbergi, including presumably topotypic samples, (12) P. sp. Tanzania 1 in the suburbs of the city of Arusha, on the eastern side of the Great Rift Valley, (13) P. sp. Tanzania 2, located in “Klein’s Camp” at the northeastern tip of Serengeti National Park, (14) P. sp. Katanga 1 in eastern Katanga Province,

DRC, and (15) P. sp. Katanga 2 at the southernmost side of the latter province. It is likely that P. sp. Mozambique 6 from northwestern Mozambique is also a unique lineage, based on its distinctive branch length.

19 20 Figure 3.1.1. Map of central, eastern and southern Africa showing the disjunct distribution of Panaspis wahlbergi (in orange). Sampled localities for the genetic analyses are shown with circles. Circle colors correspond to the clades in Figure 1.3.1. Map was modified from Branch (1998) and Spawls et al. (2002). Black circles indicate type localities of Ablepharus anselli (Kasempa, Zambia), Ablepharus moeruensis (Kilwa Island, Lake Mweru between Zambia and Katanga, DRC) and Panaspis seydeli (Lubumbashi, southeast Katanga, DRC). All of these taxa are currently considered to be synonyms of Afroablepharus seydeli (Uetz and Ho!ek, 2015).

3.2 Divergence time estimation

Results from our mitochondrial analysis suggested a basal divergence of the

Panaspis/Afroablepharus clade in the late Paleocene around 56.48 mya (51.34–61.96 mya, 95% highest posterior densities [HPD]) (Fig. 3.2.1, Appendix I). While our mitochondrial BEAST analyses yielded basal root divergences in the Eocene and Oligocene, most radiations of

Panaspis/Afroablepharus lineages occurred during the Miocene (Appendix I).

21

Figure 3.2.1. Mitochondrial (cyt b) phylogenetic tree from BEAST analyses. Open circles denote calibration points, and numbers at the base of nodes denote mean highest posterior densities (HPD).

22 The basal divergence for Panaspis/Afroablepharus recovered in our nuclear BEAST dating tree was in the late Eocene, 37.16 mya (25.29–50.37 mya, 95% HPD), with a difference of about 20 mya later than the mitochondrial results (Fig. 3.2.2, Appendix I). Our nuclear BEAST dating analysis yielded slightly younger results from the mitochondrial analysis, with basal nodes starting in the Oligocene and Miocene, and further radiations occurring during the Miocene and

Pliocene (Appendix I).

23

Figure 3.2.2. Nuclear (RAG1) phylogenetic tree from BEAST analyses. Open circles denote calibration points, and numbers at the base of nodes denote mean highest posterior densities (HPD).

24 Because some samples from our concatenated analyses failed to amplify for cyt b and

RAG1, the clades they formed are missing from our divergence time estimations (Appendix I).

Including them as missing in our data set could potentially yield problematic results (Blankers et al., 2013). Many samples are not placed in their corresponding well-supported clade from the concatenated analyses (Fig. 3.1.1), altering the overall topology of the nuclear BEAST tree. This is attributed to incomplete lineage sorting, common in phylogenetic analyses limited to nuclear genes (Brown and Twomey, 2009; McGuire et al., 2007; Morando et al., 2007; Portillo and

Greenbaum, 2014).

25

CHAPTER 4: DISCUSSION

4.1 Biogeography

Our analyses recovered strongly supported lineages that are mainly distributed in areas reaching elevations up to 1,500 m. The clades found at the eastern side of sub-Saharan Africa are situated around the Afromontane Archipelago, which consists of a series of discontinuous mountain formations along the east African coast, ranging from the southernmost tip of South

Africa to the Arabian Peninsula (Grimshaw, 2001). Although our recovered lineages are not considered Afromontane, their divergences might be explained by the irregular physiography seen along the areas where these populations occur. This pattern of microendemism has been documented in other skinks (Parham and Papenfuss, 2009), geckos (Travers et al., 2014), chameleons (Glaw et al., 2012), chelonians (Daniels et al., 2007; Petzold et al., 2014), birds

(Huseman et al., 2013) and mammals (Stoffberg et al., 2012; Taylor et al., 2011).

For example, Tanzania has two populations that are separated by the Great Rift Valley: P. sp.

Tanzania 1, located in Arusha at 1,400 m elevation, and P. sp. Tanzania 2, located at “Klein’s

Camp” in Serengeti National Park at approximately 1,500–1,800 m elevation. Both populations are located in the disjunct Southern Acacia-Commiphora Bushlands and Thickets Ecoregion, which consists of tropical and sub-tropical grasslands and savanna (Burgess et al., 2004). Similar patterns of diversification are seen in savanna-adapted snakes (Broadley, 2001b).

Mozambique harbors the greatest diversity of skink clades found in our study. The country is dominated by tropical and subtropical grasslands, savannas and shrublands, and contains a variety of hills, low plateaus and highlands (Burgess et al., 2004). The P. sp. Mozambique 1 clade resides within the Southern Miombo Woodlands, a lowland ecoregion with mainly tropical and subtropical savannas. This ecoregion is disjunct, covering the northwestern tip and central

26 area of Mozambique. A few samples also fall inside the subhumid Zambezian and Mopane

Woodlands ecoregion (Burgess et al., 2004), which occupies most of western Mozambique and is located between the latter ecoregion’s disjunct areas. Though the Zambezian and Mopane

Woodlands have scant vertebrate endemics, reptile endemism can be observed with a few strict endemics such as Lang’s worm lizard (Chirindia langi), the collared flat lizard (Platysaurus torquatus) and the Sabi quill-snouted snake (Xenocalamus sabiensis) (Burgess et al., 2004). The region possesses an outstanding conservation status because of its poor agricultural potential

(Huntley, 1978).

The distinct clades at the northeastern tip of Mozambique (P. sp. Mozambique 2, P. sp.

Mozambique 3, P. sp. Mozambique 4, and P. sp. Mozambique 6) are located in areas with different types of habitats. Cabo Delgado Province harbors the neighboring populations of P. sp.

Mozambique 2 and 4 along the northeastern coastline. The corresponding ecoregion is called the

Southern Zanzibar-Inhambane Coastal Forest Mosaic, and one sample of P. sp. Mozambique 4 falls within the Eastern Miombo Woodlands. Even though both ecoregions contain mosaics of tropical and subtropical grasslands and savannas, the coastal mosaic forests have been reported as being “biologically valuable” (Burgess et al., 2003), harboring a great variety of plant and vertebrate endemics. The P. sp. Mozambique 3 clade is located in Nampula Province, south of

Cabo Delgado. This population falls within the Eastern Miombo Ecoregion, with elevations ranging from 300–500 m. The P. sp. Mozambique 5 clade (Fig. 3.1.1) is located within the sub- humid Coastal Zambezian Ecoregion (Burgess et al. 2004), in lowland tropical savanna. The specimen that corresponds to P. sp. Mozambique 6 was collected in high elevation grassland on

Serra Jeci (1,358 m), a massif located in the Niassa Game Reserve (NGR). Given the large number of endemic reptiles described from Mozambique in recent years (Branch et al., 2014;

27 Branch and Bayliss, 2009; Branch and Tolley, 2010; Portik et al., 2013b), it should not be surprising that the country harbors a large number of cryptic species of snake-eyed skinks.

The only population from Malawi (Fig. 3.1.1) was found near Fort Lister, an area that is in a gap between the Mt. Mulanje massif and Mchese Mountain and Likabula Forest Station on the western side of the main massif. This region is dominated by montane grasslands and shublands, and represents an important center of endemism in the Afromontane Archipelago

(Burgess et al., 2004). Mount Mulanje is a relatively well-studied Afromontane isolate located in the South Malawi Montane Forest-Grassland Mosaic (Burgess et al., 2004). Rising up to 3,000 m elevation above the Phalombe Plain at the border with Mozambique, the massif includes many herpetofaunal endemics (Broadley, 2001a; Branch and Cunningham, 2006; Günther, 1893;

Loveridge, 1953) and is currently a site of important conservation concern for amphibians

(Conradie et al., 2011). Similar studies suggest this region harbors cryptic species of other taxa, including bats (Curran et al., 2012), insects (Dijkstra and Clausnitzer, 2006) and birds (Voelker et al., 2010), which resulted from the formation of sky islands. No snake-eyed skinks are currently recorded from the upper slopes of the massif (above 1,500 m altitude).

Although specimens in Namibia were collected in localities that are distant from each other (one from the Northern Namibian Escarpment [NNE] at Sesfontein, and another from the Otavi

Highlands on the Namibian central plateau), they formed a well-supported clade with minimal genetic divergence from each other. Much of Namibia comprises xeric savanna and represents a center of high reptile diversity and endemism, but many areas remain understudied (Herrmann and Branch, 2012). However, Bauer (2010) explained that even though the NNE and the Otavi

Highlands are known for having substantial biodiversity, long-term isolation and thus endemism decrease owing to the “low relief” and accessibility to surrounding areas. This might explain the

28 close relationship between the two Namibian Panaspis sp. samples. There is also a Namibian population of mole rats with a widespread distribution (Faulkes et al., 2004), which is also attributed to low relief and high accessibility in the landscape.

According to Jacobsen and Broadley (2010), A. wahlbergi can be found in a variety of habitats, from rocky outcrops to highveld grassland at altitudes ranging from sea level to 2,000 m. Our sampling suggests that A. wahlbergi occupies mostly xeric and montane shrublands and grasslands in South Africa with an elevation ranging from 10–1,300 m. The presumed distribution of A. wahlbergi has changed over time as field guides were updated through fieldwork efforts in the late 20th and early 21st centuries (Branch, 1998; Spawls et al., 2002). This species has been reported from as far as Saudi Arabia (Al-Jumaily, 1984), but misidentification is likely because of substantial habitat differences (xeric highlands instead of savanna) and the sole use of morphology in the latter study. Two unknown clades (P. sp. Limpopo and P. sp.

Gauteng, Figs. 1–2) are sympatric with populations of A. maculicollis and A. wahlbergi, respectively. Located in an area dominated by diverse habitats and high endemism (Burgess et al., 2004), they likely represent new species of Panaspis, because no other recognized species are known from that area. Makhubo et al. (2015) recently found a similar pattern in South African geckos (Afroedura) in the family Gekkonidae.

We recovered two clades residing within the Central Zambezian Miombo Woodlands

Ecoregion (P. sp. Katanga 1 and P. sp. Katanga 2) in Katanga (DRC) and northern Zambia, which contains high physiographic diversity. Southeast Katanga is dominated by various high relief areas, which contain numerous ravines, depressions and drainage systems. This region is dominated by miombo/woodland savanna (Burgess et al., 2004) and harbors various hotspots for plant and reptile endemism (Broadley and Cotterill, 2004). Plateaus in southeast Katanga are

29 believed to have formed from sands in the Plio-Pleistocene that coincided with extensive aridification processes. In an area with such geological and vegetation complexity, it is not surprising to recover unknown skink species as we did in our phylogeny (Fig. 3.1.1). The extremely close morphological resemblance between all species known from Katanga justifies the actions of earlier herpetologists, who merged Panaspis anselli, P. moeruensis, P. smithii and

P. seydeli into a single species, Afroablepharus seydeli. However, considering the extensive habitat diversity in Katanga and the large number of lineages recovered in our phylogeny, all four species could potentially be elevated to full species with additional sampling and morphological evidence.

4.2 Divergence dating

According to both dating analyses, most of the Panaspis/Afroablepharus lineages diverged between the late Oligocene and Pleistocene (Figs. 3–4). Even though similar calibration points were used, there is an approximately 20 my difference in date estimates between our mitochondrial and nuclear analyses. However, other divergence dating studies (Hugall et al.,

2007; Jones et al., 2013; Mulcahy et al., 2012) suggested that a convincing estimation could still be obtained despite similar mitochondrial/nuclear gene discrepancies in dating estimates. Our mitochondrial dating analyses yielded older calibration estimates for most of our clades.

Saturation is known to have a significant effect on mitochondrial genes because of their rapid mutation rates, resulting in overestimated divergence dates (Hugall et al., 2007;

Lukoscheck et al., 2012; Mulcahy et al., 2012). Basal branches tend to be compressed and internal nodes are pushed toward basal nodes. Nuclear genes are typically used for divergence studies above the species level, because they are slowly evolving and not prone to saturation

30 (Skinner et al., 2011; Mulcahy et al., 2012; Pyron et al., 2013). However, the branch lengths between lineages might be larger if not combined with mitochondrial loci, as suggested by

Brandley et al. (2011). In our case, basal branch lengths increased in our nuclear data set, whereas basal nodes were compressed in our mitochondrial data set. It must also be considered that we used only one nuclear locus, as compared to Mulcahy et al. (2012), who favored multiple sampling of nuclear loci. Dating estimates are vulnerable to limited sampling of loci and the sole use of mitochondrial data, which can result in differences between data sets that exceed 50 my.

Because of the unavailability of matching mitochondrial and nuclear sequences for the same samples, we presented the two analyses separately and recovered mostly congruent results for the derived lineages in our trees.

The topology of our nuclear dating tree (Fig. 3.2.2) was altered in comparison to the concatenated gene tree (Fig. 3.1.1). Some samples of P. sp. Mozambique 1 render their corresponding clade paraphyletic. Samples of A. wahlbergi and P. sp. Katanga 1 are allocated to other clades. This is likely attributable to incomplete lineage sorting of the highly conservative

RAG1 gene.

Our dating analyses suggest that the genus Panaspis first emerged in the Eocene, when savanna and grassland habitats began to appear following global cooling and the fragmentation of the pan-African forest (Tolley et al., 2011). Diversification continued from the early Miocene to the early Pliocene as cooling conditions progressed, causing the expansion of ideal habitats for

Panaspis in sub-Saharan Africa. The presence of wind-pollinated taxa and grazing vertebrates in the fossil record helped determine that main savanna development in southern Africa took place from the early Miocene to the Holocene (Jacobs, 2004). This timeframe coincides with the transition from C3 to C4 vegetation, which altered the diets of many mammalian grazers and

31 caused shifts in their distribution (Sepulchre et al., 2006). Northern and southern savanna areas increased in East Africa during the mid-Miocene, encouraging colonization by various vertebrate lineages to open habitats (Voelker et al., 2012). Transition from woodlands to grasslands in the

Miocene is also attributed to alterations in the concentration of atmospheric CO2 caused by cooling of the Indian Ocean and glacial cycles (Sepulchre et al., 2006). Further climate changes were also caused by rifting processes such as the formation of the East African Rift System in the early Oligocene and its completion in the mid-Miocene (Roberts et al., 2012). Global temperature changes during the Pliocene caused the Afrotropical forest to expand eastward to coastal Kenya and resulted in the division of the northern and southern savanna regions (Voelker et al., 2012). This loss of savanna habitat connectivity triggered diversification of major vertebrate, arid-adapted lineages, which might explain the emergence of divergent Panaspis populations from Katanga, Mozambique, South Africa and Tanzania (Fig. 3.2.2).

We found congruency between the ages of diversification of our clades and climatic and geologic events in sub-Saharan Africa. The western branch of the East African Rift System, covering northern Mozambique, formed in the late Oligocene around 25–26 mya (Roberts et al.,

2012), and its completion thereafter (~20 mya) coincides with the radiations in our Mozambique clades (Figs 3.2.1–3.2.2). Tiercelin and Lezzar (2002) suggested that the Eastern Arc Mountains and Southern Highlands of Tanzania arose during the late Miocene. Climatic shifts also took place during that time and encouraged the development of forest refugia in the region (Menegon et al., 2014). The sister clades from Arusha and Serengeti shared a common ancestor during this period (Figs. 3.2.1–3.2.2), and these climatic and orogenic changes likely promoted their allopatric speciation. Fossil records presented by Rage (2003) supported the production of rich reptile faunas during the Miocene in Namibia. There is congruency between our speciation

32 patterns and aridification processes in that epoch, and the dates in our BEAST trees concur (Figs.

3.2.1–3.2.2). Tolley et al. (2008) described southwestern Africa as a “cradle of diversity” for species that survived the transition from C3 to C4 plant habitats. The extinction of C3-dependent species implies that while not all species survived this transition, the remaining ones had the opportunity to diversify, thus creating a biodiversity hotspot. The presence of four sympatric populations in South Africa (A. maculicollis, A. wahlbergi, P. sp. Gauteng and P. sp. Limpopo) can be explained by this concept.

Similar patterns and timing of diversification have been demonstrated in other vertebrate groups with non-forest distributions, including African clawed frogs (Furman et al., 2015), cobras (Trape et al., 2009) and lizards (Diedericks and Daniels, 2014; Makoha et al., 2007), which have all been shown to form complexes of divergent populations correlated with the expansion of C4 grasslands during the Miocene. Subsequent aridification in the Pliocene and

Pleistocene likely explain the more recent cladogenic events in our analyses (Figs. 3.2.1–3.2.2), which are similar to patterns in lions (Barnett et al., 2014; Bertola et al., 2011), mole rats

(Faulkes et al., 2004) and ungulates (Lorenzen et al., 2012).

4.3 Taxonomy and species limits

Greer (1974) erected Afroablepharus based on discrete morphological differences—the frontal scale being in contact with one supraocular, and the ablepharine eye condition. He restricted Panaspis to skinks with smooth body scales and terrestrial to fossorial habits. Greer’s

(1974) only specific characteristics for diagnosing Panapsis were having the frontal scale in contact with two supraoculars, and a pre-ablepharine eye or lower mobile eyelids, the latter only applicable to P. breviceps and P. togoensis. All examined vouchers from our study are consistent

33 with Greer’s (1974) explicit characteristics reserved for Afroablepharus. Schmitz et al. (2005) used mitochondrial data and broad sampling from Panaspis sensu lato to support the recognition of Afroablepharus as a full genus. However, they suggested an in depth assessment of Panaspis sensu stricto, because discrepancies exist in the ecology of some of its species (e.g., P. breviceps is a lowland rainforest species). Furthermore, DNA sequences of P. cabindae (the type species) were not included, thus restricting taxonomic conclusions of the latter study. Currently recognized species in Afroablepharus, including the type species A. wahlbergi, were nested within our well-supported Panaspis clade (type species P. cabindae) from Fig. 3.1.1, rendering

Afroablepharus polyphyletic. This finding warrants the synonymization of that Afroablepharus

(Greer, 1974) with Panaspis (Cope, 1868), which has priority over the former genus. The species

P. breviceps and P. togoensis formed reciprocally monophyletic clades with relatively long branch lengths (Fig. 3.1.1), thus refuting previous herpetologists’ ideas that considered P. togoensis a subspecies of P. breviceps (Hoogmoed, 1980; Loveridge, 1952). Because these two species are also highly distinct from Panaspis sensu stricto, their position as snake-eyed skinks must be reassessed (Fig. 3.1.1) in a future study. Excluding the two latter species, our molecular and morphological analyses confirm that Panaspis should accommodate savanna skinks with pre-ablepharine and ablepharine eyes, as Broadley (1989) suggested.

We recovered strong support for the reciprocal monophyly of at least 18 lineages of

Panaspis, most of which are likely to be new species. Prior to this study, herpetologists believed that Panaspis wahlbergi inhabited the enormous geographic area shown in orange in Fig. 1, and that the disjunct distribution could be explained by gaps in sampling. However, our phylogeny of

P. wahlbergi samples demonstrated that the species is included in a complex of at least 18 cryptic lineages that are morphologically similar, but distinct genetically (Fig. 3.1.1).

34 According to Spawls et al. (2002), P. wahlbergi is presumed to occupy a large area along the eastern coast of Tanzania and a small, disjunct population occurs at the southwestern tip of

Lake Tanganyika (Fig. 1.3.1). However, it is likely that P. sp. Tanzania 1 corresponds to

Panaspis megalurus, known from “the mid-altitude central plains of Tanzania, north and northwest of Dodoma” (Spawls et al., 2002). The suggested range for P. megalurus extends throughout the ecoregion it is found in, from Arusha southward to Dodoma. This range coincides with our samples from Arusha (Fig. 3.1.1). The type locality Kinjanganja in “Turu”, as written by Nieden (1913), could not be pinpointed with accuracy (only latitude coordinates were provided in the original description), but it is believed to be located in central Tanzania, close to

Dodoma (Uetz and Ho!ek, 2015) within the presumed range of this species.

Exploration of the NGR has resulted in the description of new species and identification of reptile and amphibian taxa with unresolved taxonomic status (Branch et al., 2005; Portik et al.,

2013a). Large areas of Mozambique remain largely unexplored because of inaccessibility in the

Lichinga Plateau where Serra Jeci is situated, but the NGR is known to have the highest reptile diversity in Mozambique (Branch et al., 2005). Our data suggest that multiple new species of

Panaspis are likely to occur in Mozambique. To date, there are thorough vertebrate biodiversity assessments only from the region south of the Zambezi River (Schneider et al., 2005), thus rendering the northern areas of Mozambique severely understudied. Aside from the effects of political turmoil and loss of infrastructure in southern and central Mozambique, the northern areas are so remote and underdeveloped they remain challenging for herpetological surveys

(Branch et al., 2005; Branch and Bayliss, 2009).

Our results clarified the distribution of topotypic P. wahlbergi and P. maculicollis in

South Africa (Figs. 1.3.1–3.1.1). The southern African species P. wahlbergi and P. maculicollis

35 had been previously reported from Namibia (Bauer et al., 1993; Branch, 1998; Herrmann and

Branch, 2012). However, it is unlikely that these Namibian populations are conspecific with P. wahlbergi or P. maculicollis, because the ecoregions they inhabit are completely different, and our samples from Namibia were genetically distinct (Fig. 3.1.1). Namibia is mainly dominated by arid ecoregions, whereas South Africa contains tropical and subtropical savannas.

Nonetheless, both areas share a portion of the Kalahari ecoregion, thus having similar habitats.

Further sampling is required to document the full distribution of P. sp. Namibia and describe it as a new species.

Based on distinctive morphology and proximity to type localities, we matched three lineages of our phylogeny to known species: P. wahlbergi (Smith, 1849), P. maculicollis

Jacobsen and Broadley, 2010, and P. cabindae Bocage, 1866. Panaspis wahlbergi was described from Natal, South Africa, and P. maculicollis from Klein Tsipishe, at the northeastern tip of the country. The distribution of our P. maculicollis samples suggest the species is sympatric with P. wahlbergi, because they were collected in nearby localities (Fig. 1.3.1). There are two unknown lineages of Panaspis (Panaspis sp. Gauteng and Panaspis sp. Limpopo) located in the vicinity of

P. maculicollis and P. wahlbergi. Based on our phylogenetic analyses, P. wahlbergi has a potentially large distribution that has yet to be thoroughly explored, because large gaps in sampling exist in our study (Fig. 1.3.1). The type species P. cabindae was described from Bas-

Congo Province (DRC), south of the Congo River (Bocage, 1866). Specimens from various locations in Angola and one in southwestern DRC were nested in a well-supported clade belonging to P. cabindae, demonstrating the species has a widespread distribution in south- central Africa.

36 Several snake-eyed skink species from sub-Saharan Africa lack molecular sampling and are poorly known in general. Two of these species were described from the Ruwenzori massif between Uganda and DRC more than half a century ago: Panaspis helleri (Loveridge, 1932) at

2,895 m (DRC) and Panaspis burgeoni (de Witte, 1933) at 2,073 m (DRC). The Ethiopian species, Panaspis tancredi, should retain its full species status, but extensive sampling is required to confirm its distribution, because few specimens have been found (Boulenger, 1909;

Largen and Spawls, 2006). The West African members of Panaspis, P. breviceps (Peters, 1873),

P. togoensis and P. nimbensis (Angel, 1944), need to be examined in greater detail, because they all have lower mobile eyelids. Considering remaining taxa that were formerly members of the synonymized genus Afroablepharus, P. wilsoni is only known from Sudan and P. duruarum resides in Cameroon, whereas P. africanus and P. annobonensis are located on volcanic islands of the Gulf of Guinea (Uetz and Ho!ek, 2015).

4.4 Conservation

African savannas cover most of the central and southern parts of the continent (Sodhi et al., 2007). They harbor the world’s greatest diversity of ungulates and therefore a variety of predators. Termites are also abundant and contribute to soil fertility and serve as a principal food source for many semi-fossorial reptiles. About two fifths of land in Africa is covered by savannas, and most of that land is currently used for livestock farming to sustain local populations (Hassler et al., 2010; Sodhi et al., 2007). Savannas are constantly exposed to degradation because of poor farming management, uncontrolled fires and mining, threatening biodiversity in many African countries and hotspots including the Niassa Game Reserve, Mt.

Mulanje Biosphere Reserve, Quirimbas National Park of coastal northeastern Mozambique, the

37 xeric savannas of Namibia, and the largely unprotected Katanga miombo savannas (Herrmann and Branch, 2012; Sodhi et al., 2007). However, it has been shown that many species of lizards in savanna and semi-aquatic habitats are resilient after fires (Andersen et al., 2013; Costa et al.,

2013; Gorissen et al., 2015), and other anthropogenic disturbances (Smart et al., 2005). Several specimens in our study were found in disturbed areas, such as mining concessions (Appendix I) and an outhouse in the Bombo-Lumene Game Park (EG, pers. comm.). While it is likely that some of these species occur in relatively small populations, future studies are needed to determine whether Panaspis/Afroablepharus should be given a threatened status.

38

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52 APPENDIX I

Table 1. Field numbers and localities for specimens used in genetic analyses. DRC = Democratic Republic of Congo, E = east, Moz = Mozambique, N = north, NW = northwest, SW = southwest, S = south, SA = South Africa. Species Field Number Collection Number Locality 16S cyt b PDC RAG1 Cordylus marunguensis EBG 2993 — Pepa, Katanga, DRC — — — — Trachylepis megalura EBG 1409 — Lwiro, South Kivu, DRC — — — — Trachylepis striata EBG 1407 — Lwiro, South Kivu, DRC — — — — Panaspis cabindae 804 PEM R20256 Soyo, NW Angola — — — — Panaspis cabindae PM 050 — Bas-Congo, DRC — — — — Panaspis cabindae PM 049 — Bas-Congo, DRC — — — — Panaspis cabindae 810 WC1820 Riverine Forest, Bengo, Angola — — — — Panaspis cabindae ANG21 WC 1006 Lagoa Carumbo, Angola — — — — Panaspis cabindae ELI 1722 — Bombo-Lumene Reserve, Kinshasa, DRC — — — — Panaspis cabindae ANL 52 MTD 48612 Kimpa Vita Uni Campus, Uíge, N Angola — — — — Panaspis cabindae MBUR 2128 665 S Leba Pass, Huila District, SW Angola — — — — Panaspis sp. Ethiopia TJC 264 — Oromia, western Ethiopia — — — — Panaspis sp. Limpopo MCZA 27176 — Hoedspruit, Limpopo, SA — — — — Panaspis sp. Limpopo MCZA 27177 CAS 248791 Hoedspruit, Limpopo, SA — — — — Panaspis sp. Namibia AMB 7634 MCZ R-183767 Sesfontein, Namibia — — — — Panaspis sp. Namibia — 567 Otavi, Namibia — — — — Panaspis maculicollis MBUR 02843 uncatalogued Phalaborwa, Limpopo, SA — — — — Panaspis maculicollis MBUR 02848 uncatalogued Phalaborwa, Limpopo, SA — — — — Panaspis maculicollis MCZF 38848 CAS 234188 Limpopo Province, SA — — — — Panaspis maculicollis MCZF 38790 CAS 234135 Limpopo Province, SA — — — — Panaspis maculicollis MCZF 38733 CAS 234099 Limpopo Province, SA — — — — Panaspis sp. Mozambique 1 WC 1251 PEMR 20561 Ecofarm, Chemba, Moz — — — — Panaspis sp. Mozambique 1 WC 1249 no voucher Ecofarm, Chemba, Moz — — — —

53 Panaspis sp. Mozambique 1 WC 1169 PEMR 20565 Boabab Ore Mine, Masamba, Moz — — — — Panaspis sp. Mozambique 1 WC 1186 PEMR 20566 Boabab Ore Mine, Masamba, Moz — — — — Panaspis sp. Mozambique 1 SVN 693 — Gorongosa National Park, Moz — — — — Panaspis sp. Mozambique 1 886 CAP 22 Ruoni Hill S, Tete Province, Moz — — — — Panaspis sp. Mozambique 2 WC 1358 uncatalogued Quiterajo, Cabo Delgado, Moz — — — — Panaspis sp. Mozambique 2 ENI 038 uncatalogued Mocimboa da Praia, Cabo Delgado, Moz — — — — Panaspis sp. Malawi 568 PEMR 20247 Fort Lister, Mt. Mulanje, Malawi — — — — Panaspis sp. Malawi 570 PEMR 9752 Likabula Station, Mt. Mulanje, Malawi — — — — Panaspis sp. Mozambique 3 WC 1051 no voucher NW of Rapale, Nampula, Moz — — — — Panaspis sp. Mozambique 3 WC 1067 PEM R20557 E of Ribuae, Nampula, Moz — — — — Panaspis sp. Mozambique 3 WC 1133 no voucher NW of Mecuburi, Nampula, Moz — — — — Panaspis sp. Mozambique 3 WC 1161 PEMR 20558 Rapale, Nampula, Moz — — — — Panaspis sp. Mozambique 4 855 Syran 20 Syran graphite mine, Balama, Moz — — — — Panaspis sp. Mozambique 4 856 Syran 21 Syran graphite mine, Balama, Moz — — — — Panaspis sp. Mozambique 4 WC 1317 uncatalogued Pemba, Cabo Delgado, Moz — — — — Panaspis sp. Mozambique 4 WC 1404 uncatalogued Pemba, Cabo Delgado, Moz — — — — Panaspis sp. Mozambique 4 ENI 037 uncatalogued Quirimbas National Park, Moz — — — — Panaspis sp. Mozambique 5 SVN 742 NMB R10286 Beira, Mozambique — — — — Panaspis sp. Gauteng WRB 952 WC 0214 Doornfontein, Gauteng, SA — — — — Panaspis sp. Gauteng WRB 953 WC 2721 Doornkop, Gauteng, SA — — — — Panaspis wahlbergi DMP 127 MVZ 266147 Inhambane, Mozambique — — — — Panaspis wahlbergi — TM 84299 Groblersdal, Limpopo, SA — — — — Panaspis wahlbergi MCZF 38852 CAS 234194 Limpopo Province, SA — — — — Panaspis wahlbergi AMB 8279 R 184432 Limpopo Province, SA — — — — Panaspis wahlbergi AMB 8293 R 184443 Limpopo Province, SA — — — — Panaspis wahlbergi MCZF 38868 CAS 234209 Limpopo Province, SA — — — — Panaspis wahlbergi TJH 3253 95588 Kimberley, Northern Cape, SA — — — — Panaspis wahlbergi TJH 3213 95563 Kimberley, Northern Cape, SA — — — — Panaspis sp. Mozambique 6 DMP 187 MVZ 266148 Niassa, Moz — — — —

54 Panaspis sp. Tanzania 1 WRB 0021 — Arusha, Tanzania — — — — Panaspis sp. Tanzania 1 WRB 0026 — Arusha, Tanzania — — — — Panaspis sp. Tanzania 2 572 No voucher Klein's Camp, Serengeti, Tanzania — — — — Panaspis sp. Tanzania 2 573 No voucher Klein's Camp, Serengeti, Tanzania — — — — Panaspis sp. Katanga 1 ELI 294 — Katanga, DRC — — — — Panaspis sp. Katanga 1 ELI 295 — Katanga, DRC — — — — Panaspis sp. Katanga 2 575 PEM R17454 Kalakundi Copper Mine, S Katanga, DRC — — — — Panaspis sp. Katanga 2 576 PEM R17455 Kalakundi Copper Mine, S Katanga, DRC — — — — Panaspis sp. Katanga 2 JHK 26 uncatalogued Kisanfu Camp, Katanga, DRC — — — — Panaspis sp. Katanga 2 WRB 0047 PEM R20327 Fungurume Camp, Katanga, DRC — — — — Panaspis sp. Katanga 2 WRBNimb083 — NW Zambia — — — — Afroablepharus africanus — uncatalogued Príncipe, Gulf of Guinea — — — — Afroablepharus africanus — — Montalegre, Príncipe, Gulf of Guinea EU164477 — — — Afroablepharus africanus — uncatalogued Príncipe, Gulf of Guinea AY308286 — — — Afroablepharus annobonensis — — Annobon, Gulf of Guinea EU164494 — — — Afroablepharus annobonensis — — Annobon, Gulf of Guinea EU164488 — — — Mochlus afer — No voucher Kenya — — — — Lacertaspis reichenowi — — — AY308235 — — — Lacertaspis rohdei — ZFMK 75382 Mt. Nlonako, Cameroon — — — — Lacertaspis chriswildi — ZFMK 75735 Tchabal Mbabo, Cameroon — — — — Lacertaspis gemmiventris RCD 13251 CAS 207854 Bioko Island, Equatorial Guinea — — — — Lacertaspis gemmiventris RCD 13255 CAS 207858 Bioko Island, Equatorial Guinea — — — — Leptosiaphos koutoui — MNHN 2001.0697 Meiganga, Adamaoua Plateau, Cameroon — — — — Leptosiaphos blochmanni — ELI 1610 Bichaka, South Kivu, DRC — — — — Leptosiaphos meleagris — ELI 2844 Ruwenzori National Park, Uganda — — — — Leptosiaphos sp. — ZFMK 69552 Mt. Nlonako, Cameroon — — — — Leptosiaphos sp. — uncatalogued Mt. Nlonako, Cameroon — — — — Leptosiaphos sp. — ZFMK 75381 Mt. Nlonako, Cameroon — — — — Panaspis breviceps — ELI 558 Byonga, South Kivu, DRC — — — —

55 Panaspis breviceps — MM 106 Mawne, Cameroon — — — — Panaspis breviceps — MM105 Mawne, Cameroon — — — — Panaspis breviceps — ZFMK 75380 Mt. Nlonako, Cameroon — — — — Panaspis breviceps — uncatalogued Cameroon — — — — Panaspis togoensis — ZFMK 42212 — — — — — Panaspis togoensis 2426 MVZ 249793 Kyabobo National Park, Ghana — — — — Panaspis togoensis DCB 34707 — Gashaka Gumti National Park, Nigeria — — — — Panaspis togoensis TJH 2561 94519 W National Park, Alibori, Benin — — — — Panaspis togoensis TJH 2629 94557 Dogo Forest, Benin — — — — Panaspis togoensis TJH 2600 94544 W National Park, Alibori, Benin — — — — Karusasaurus polyzonus — ABTC03038 — — — — EU366444 Namazonurus namaquensis AMB 6848 CAS 223963 Karas Region, Namibia — EU116507 — — Namazonurus namaquensis — — — — AY217796 — — Smaug mossambicus — — — — DQ090879 — — Trachylepis varia — — — — — — GU931670 Trachylepis punctulata — — — — — — GU931678 Trachylepis punctulata — — — — — — JX568135 Trachylepis punctulata — — — — — — JX568133 Trachylepis variegata — — — — — — JX568152 Trachylepis variegata — — — — — — JX568151 Trachylepis variegata — — — — — — JX568148 Trachylepis variegata — — — — — — GU931676 Trachylepis variegata — — — — DQ239179 — — Trachylepis sulcata — — — — — — GU931665 Trachylepis sulcata — — — — — — GU931664 Trachylepis sulcata — — — — — — GU931663 Trachylepis vittata — — — — EU443142 — — Trachylepis perrotetii — — — — DQ239146 — — Trachylepis perrotetii — — — — DQ239145 — —

56 Trachylepis perrotetii — — — — DQ239142 — — Xantusia vigilis — — — — EU116665 — — Xantusia vigilis — — — — EU116664 — — Xantusia vigilis — — — — EU116663 — — Scelotes gronovii — — — — JN561448 — — Scelotes gronovii — — — — JN561438 — — Scelotes kasneri — — — — JN561423 — — Scelotes kasneri — — — — JN561421 — — Scelotes sexilineatus — — — — JN561420 — — Scelotes bipes — — — — JN561411 — — Tiliqua gigas — — — — AY217811 — — Tiliqua rugosa — — — — — — EF534815 Plestiodon fasciatus — — — — AY217818 — — Plestiodon fasciatus — — — — DQ241673 — — Plestiodon fasciatus — — — — DQ241668 — — Feylinia grandisquamis — — — — AY217798 — — Feylinia polylepis — — — — FJ972225 — — Feylinia polylepis — — — — EU164536 — — Trachylepis homalocephala — — — — — — GU931615 Mabuya dominicana — — — — JN227561 — — Trachylepis quinquetaeniata — — — — EU443143 — — Mabuya bistriata — — — — DQ239187 — — Mabuya frenata — — — — DQ239173 — —

57 Estimated dates of divergence and highest posterior densities (HPD) from our mitochondrial BEAST analyses of clades in the Panaspis/Afroablepharus complex Node Date Age 95% HPD Panaspis/Afroablepharus 56.48 Paleocene 51.34–61.96 P. breviceps, P. togoensis 36.03 Eocene 29.09–42.78 P. cf. Ethiopia, P. sp. Limpopo, P. sp. Namibia, A. maculicollis, P. sp. Mozambique 1 42.83 Eocene 37.15–48.69 P. sp. Limpopo, P. sp. Namibia 21.77 Miocene 16.93–26.77 A. maculicollis, P. sp. Mozambique 1 18.05 Miocene 13.7–22.2 A. africanus, P. cabindae 35.14 Eocene 28.31–42.02 P. cabindae 16.15 Miocene 11.92–20.52 P. sp. Mozambique 4, P. sp. Malawi, P. sp. Mozambique 3 17.47 Miocene 13.87–21.3 P. sp. Malawi, P. sp. Mozambique 3 13.76 Miocene 10.09–17.71 P. sp. Gauteng, P. sp. Mozambique 5, A. wahlbergi 20.58 Miocene 16.57–24.7 P. sp. Mozambique 5, A. wahlbergi 15.31 Miocene 11.96–18.77 P. sp. Mozambique 6, P. sp. Tanzania 1, P. sp. Tanzania 2, P. sp. Katanga 1, P. sp. Katanga 2 25.69 Oligocene 21.62–29.68 P. sp. Mozambique 6, P. sp. Tanzania 1, P. sp. Tanzania 2 19.43 Miocene 15.21–23.68 P. sp. Katanga 1, P. sp. Katanga 2 19.25 Miocene 14.76–23.61

58 Estimated dates of divergence and highest posterior densities (HPD) from our nuclear BEAST analyses of clades in the Panaspis/Afroablepharus complex Node Date Age 95% HPD Panaspis/Afroablepharus 37.16 Eocene 25.29–50.37 P. cabindae 27.27 Oligocene 17.96–38.05 P. sp. Namibia, P. sp. Limpopo, P. togoensis, P. sp. Mozambique 1, A. maculicollis 11.82 Miocene 6.25–17.49 P. sp. Namibia, P. sp. Limpopo, P. sp. togoensis 8.58 Miocene 4.15–13.45 P. sp. Mozambique 1, A. maculicollis 6.22 Miocene 2.01–10.63 P. sp. Malawi, P. sp. Mozambique 4 5.95 Miocene 2.04–10.42 P. sp. Tanzania 1, P. sp. Katanga 1, P. sp. Katanga 2 4.71 Pliocene 1.91–7.67 P. sp. Katanga 1, P. sp. Katanga 2 3.16 Pliocene 0.97–5.49 P. sp. Gauteng, P. sp. Mozambique 5, A. wahlbergi 2.98 Pliocene 0.69–5.62 P. sp. Mozambique 5, A. wahlbergi 1.72 Pliocene 0.21–3.72 A. wahlbergi 2.54 Pliocene 0.38–5.18 A. wahlbergi, P. sp. Tanzania 1, P. sp. Tanzania 2 6.05 Miocene 2.97–9.35 P. sp. Katanga 2, P. sp. Tanzania 1, P. sp. Tanzania 2 5.22 Miocene 1.83–8.41

59 VITA

Maria Fernanda Medina began her college career in 2009, completing a B.S. in

Biomedical Sciences at The University of Texas at El Paso (UTEP) in May 2012. During her time at UTEP, she participated in several career-oriented organizations and field biology courses.

During the spring of 2010, Maria traveled to Costa Rica’s Osa Peninsula to participate in a conservation project that involved monitoring dolphin populations and measuring poison dart frogs. She also became the Veterinary Representative for the Medical Professions Organization at UTEP and an active member for the American Society for Microbiology. Her brief exposure to field work inspired her to begin a research project involving reptiles and amphibians. In spring of 2011, Maria became a Special Problems student with Dr. Eli Greenbaum and continually applied to research programs, such as the UTEP College Office of Undergraduate Research

Initiatives (COURI), and the UTEP Research Initiatives for Scientific Enhancement (RISE).

Maria began her graduate degree at UTEP in August 2012. During this time, she received a scholarship from the National Science Foundation (NSF) to support her research and mentor an undergraduate student. Maria has also been a teaching assistant for Organismal Biology (BIOL

1108) and Birds and Mammals (BIOL 4478). She is a member of the Society for the Study of

Amphibians and Reptiles, The Herpetologist’s League and the American Society of

Ichthyologists and Herpetologists. She presented her graduate research at the World Congress of

Herpetology meeting in Vancouver, Canada (August 2012), the Joint Meeting of Ichthyologists and Herpetologists in Albuquerque, NM (July 2013) and Chattanooga, TN (August 2014) and the Society for Molecular Biology and Evolution meeting in San Juan, Puerto Rico in June 2014.

She plans to pursue a combined Doctor of Veterinary Medicine and doctoral degree

(DVM/Ph.D.) to contribute to herpetology as well as the growing herpetocultural community.

60 Permanent address: 1337 Vista de Oro Dr.

El Paso, TX 79935

This thesis/dissertation was typed by Maria Fernanda Medina

61