A Phylogeny of Genera Spirobolomyia and (Diptera: Sarcophagidae)

A thesis submitted to the

Graduate School

Of the University of Cincinnati

In partial fulfillment of the requirements for the degree of

Master of Science

In the Department of Biological Sciences of the McMicken College of Arts and Sciences By

Stephanie J. Gierek

B.S., University of Cincinnati May 2014

Committee chair: Ronald W. DeBry, Ph.D. Abstract

Blaesoxipha and Spirobolomyia are two genera from the large family Sarcophagidae, more commonly known as flesh . These two genera are parasitoids to a wide variety of other . When Spirobolomyia was first described by Townsend in 1917, it was a new genus with only one species. In 1965, Downes synonymized Spirobolomyia with Blaesoxipha, giving it subgeneric status, and included 3 new species. In 1996, Pape added another species and removed Spirobolomyia from Blaesoxipha returning it to genus status. In this study, I infer a phylogeny using the cytochrome oxidase I (COI) and NADH dehydrogenase (ND4) genes from the ’s mitochondrial DNA (mtDNA) in hopes of determining if Spirobolomyia should remain a genus or be reduced to subgeneric status under Blaesoxipha. The phylogeny infers strong support for monophly of Spirobolomyia. However due to no support from the Maximum

Likelihood analysis but support from the Bayesian Inference, of the monophyly of Blaesoxipha, more data is needed to determine the placement of Spirobolomyia.

ii

iii Acknowledgements

First, I would like to thank my ever-patient husband. When we agreed I would start this journey four years ago, I promised him it would only take two years. Now here we are, four years later. And even though it took me twice as long, he as was always supportive. Whenever I doubted whether I could do this, he was always there to remind me I could do it. Josh, this is not only my victory but yours too. We did this together because without your support and constant confidence boots, I would have never made it this far.

I would also like to thank Dr. Ron DeBry. Little did he know when he talked me into this journey, he would have to put up with me and my life happenings for four long years. Had it not been for him I would never have thought graduate school was an option. Ron, thank you for pushing me to do this. And although it took longer than we both anticipated, I am thankful you had the faith in me to finish.

I want to say a special a thanks to Dr. Evan Wong. Without his vote of confidence, I would have never become a part of the lab. He taught me everything I needed to know when it came to lab, even the quirky little tricks it would sometime take to make something work. Evan, I was always fascinated by

iv your drive to accomplish more. I know you will accomplish great things as Dr2

Evan Wong.

I would also like to thank Dr. Greg Dahlem and Dr. Eric Tepe. I truly appreciate you taking the time from your busy schedules to be a part of my committee. Also, thank you for putting up with me and my last-minute neediness.

I would like to give a quick thank you to my new PI, Dr. Latha Satish. You welcomed me into your lab and have given me amazing support while trying to finish up my thesis.

v Table of Contents

Abstract……………………………………………………………………………………………………………… ii

Acknowledgements…………………………………………………………………….….……….………… iv

Table of Contents………………………………………………………………………….…………………. vi

List of Tables and Figures…………………………………………………………………….…………. vii

Introduction……………………………………………………………………………………………...……… 1

Materials & Methods…………………………….………………………………………………….………. 4

Results……………………………………………………………………………………….……….……………… 7

Discussion………………………………………………………………………….……………….…...……… 11

Conclusion ………………………………………………………………………………………………………. 13

Tables…………………………………………………………………………………………………………………14

References…………………………………………….………………………………………………………….20

vi Tables and Figures

Figure 1. The Bayesian phylogeny of Blaesoxipha and Spirobolomyia inferred using

mtDNA genes COI and ND4 ……………………………….………………………………….10

Table 1. Specimen Identification: Genus, subgenus, species and voucher ……….14

Table 2. List of primers used in this study…………………………………………………………19

vii

Introduction

Blaesoxipha and Spirobolomyia are two genera from the large family Sarcophagidae, more commonly known as flesh flies. These two genera are parasitoids to a wide variety of other arthropods. Blaesoxipha primarily parasitizes grasshoppers and beetles, while

Spirobolomyia preys on millipedes. Since these genera were described, entomologist have disagreed over whether these two groups are congeners or whether they should be recognized as two separate genera.

Blaesoxipha was established by Loew in 1861. He discovered a female Blaesoxipha attempting to deposit her larva on a grasshopper using a long sword-like ovipositor. The sword- like ovipositor lead to the name Blaesoxipha, blaisos meaning “Bandy-legged, bent” and xiphos meaning “sword, saber” (Pape 1994). Blaesoxipha is one of the more species-rich taxa in the family of Sarcophagidae. It has 242 described species currently organized into ten subgenera

(Pape 1994). The largest of the ten subgenera is Blaesoxipha sensu stricto. It encompasses 75 species, all of which have an old-world distribution except for two species, Blaesoxipha

(Blaesoxipha) atlantis and Blaesoxipha (Blaesoxipha) opifera, which have a New World distribution. The Subgenus Acanthodotheca is nearly as large, with 71 described species attributed to it. The species in Acanthodotheca are primarily beetle parasitoids. It has an exclusive New World distribution. The subgenera Abapa (eight species) and Aldrichisca (three species) are restricted to the Antilles. Subgenera Acridiophaga (containing 11 species) and

Servaisia (32 species) are Neotropic, Nearctic and Palearctic in distribution. Tephromyia is the only subgenus that is found only in the Palearctic region. It contains 21 species. Subgenera

1 Gigantotheca (17 species) and Kellymyia (three species) are both distributed among the New

World. The subgenus Speciosia contains only one species. Blaesoxipha speciosa, originally designated Fletcherimyia speciosa by Downes (1947), was placed in a subgenus alone by

Roeback in 1954 (Pape 1994).

A vast majority of Blaesoxipha are parasitoids of grasshoppers and darkling beetles.

However, there are species that are parasitoids to a large range of other taxa (Pape

1994). Blaesoxipha are ovoviviparous; meaning they incubate their eggs within a uterus or pouch. The eggs hatch within the uterus or pouch and first instar larvae are expelled through the larvapositor (Allen and Pape 1994). Blaesoxipha deposit or insert their larva into another insect species. Blaesoxipha have been observed depositing larva on to a host in two ways.

Females will stalk their prey from a stationary position above their prey. They will then dart out and deposit a single larva either directly on the prey or even directly within the prey via the genital-anal orifice (Leonide 1964), the buccal cavity (Salvin 1958), or directly into the haemocoele through the cuticle or between the soft segments of the cuticle (Middlekauf 1958;

Leonide 1967). Other species of Blaesoxipha will bend the abdomen forward and position their larvapositor between their legs and expel the larva forcefully onto the host. Larvae will quickly penetrate the host (Leonide and Leonide 1986). Larvae will remain inside the host haemocoele for 4-10 days. The host will usually survive while larvae develop within their haemocoele, and if large enough, may survive the exiting of the larva (Greathead 1963). After leaving the host, larvae burrow into the soil to pupate for 1-4 weeks (Crouzel and Slavin 1961).

Spirobolomyia was originally described as a genus by Townsend (1917). Spirobolomyia has only five known species (Pape, 1994). Three of the five species of Spirobolomyia were

2 described before Townsend described the genus. All three were first identified as a part of the genus . Spirobolomyia basalis was originally described by Walker in 1853 and designated Sarcophaga basalis. Sarcophaga pallipes, by original designation, was synonymized with Spirobolomyia basalis by Downes in 1965. When Aldrich (1916) described Sarcophaga pallipes, he was unable to determine the species as the holotype is female, making it difficult to identify to species with any certainty (Pape 1990). In 1916, Aldrich described a species he named Sarcophaga deceptiva in which he described to be similar Sarcophaga singularis with only minor differences. This species later was discovered to be Spirobolomyia basalis.

Spirobolomyia flavipalpis was originally identified as Sarcophaga flavipalpis by Aldrich in 1916.

Hall (1927) and Roback (1954) described two species (Sarcophaga flavipalpus and Sarcophaga flavipes) that were discovered to be Spirobolomyia flavipalpis. Hall (1927) discovered a species he named Sarcophaga cingularis that was also later identified to be Spirobolomyia singularis.

Hall (1927) also described a species he named Sarcophaga ohioensis. These four species were included in Spirobolomyia by Downes in 1965 when he placed Spirobolomyia as a subgenus of

Blaesoxipha. In 1990, Pape described the fifth species of Spirobolomyia, Spirobolomyia latissima. All five species have a Nearctic distribution with Spirobolomyia ohioensis and

Spirobolomyia latissimi expanding to the Neotropic region.

Spirobolomyia lay their larvae on dead and/or injured millipedes. Beyond the identification and knowledge of its parasitic behavior, there is little knowledge of the natural history of this genus. Spirobolomyia and Blaesoxipha share several identifying morphological characters. Both genera have a male recurved cercal prong with short spines, presence of a midfemoral comb and female fused sternites 6-8 (Pape 1990). The character that led Lopes to

3 his belief that Spirobolomyia and Blaesoxipha “have no relationship on the genus level” was the identification of the tubular functional (sperm conducting) lateral styli of Spirobolomyia which contrasts with the flat plate-like and non-sperm conducting lateral styli that are characteristic of Blaesoxipha.

Townsend (1971) described Spirobolomyia and transferred Sarcophaga singularis into his new genus. Downes (1965) synonymized Spirobolomyia with Blaesoxipha, reducing it to subgeneric status. In 1975, Lopes claimed, “these two genera had no relationship on the genus level.” In 1996, Pape removed Spirobolomyia from Blaesoxipha and returned it to full genus rank.

Before the use of phylogenetics, Spirobolomyia has been described as a genus, reduced to a subgenus and reclassified as a genus. The goal of this study was to determine whether mitochondrial DNA (mtDNA) will support a two-genus phylogeny or suggest synonymizing the genera. A phylogeny was inferred using the cytochrome oxidase I (COI) and NADH dehydrogenase (ND4) genes from the fly’s mitochondrial DNA (mtDNA). Two phylogenies were inferred using Bayesian Inference and Maximum Likelihood. I expect Spirobolomyia to be monophyletic. If a monophyletic Spirobolomyia is found to be nested within Blaesoxipha, that would support reclassifying Spirobolomyia as a subgenus of Blaesoxipha. If Spirobolomyia is found to be distinct from Blaesoxipha, that would support retention of the current classification as a distinct genus.

Methods

Collection/Extraction

4 Individuals for this study were collected by Trevor Stamper, Greg Dahlem, Evan Wong and Ronald DeBry (Stamper 2008; Wong 2015). All individuals were identified by Greg Dahlem, pinned as vouchers, and 3 legs were removed and placed in ethanol. The removed legs were stored at -80°C until ready for extraction.

For extraction, 1-2 legs were removed from the previously frozen specimen and cut into fourths. DNA was extracted using a Qiagen DNeasy kit (Cat ID 69504). Except for the final step, in which DNA grade water was used in place of AE elution buffer, the protocol for blood/tissue extraction provided by Qiagen was followed. The extracted DNA samples were then stored at

0C until used for amplification.

Amplification/Sequencing

Mitochondrial DNA was amplified with primers designed to amplify regions that encode portions of the proteins Cytochrome Oxidase I and NADH dehydrogenase (Table 2). Small scale amplifications were performed first to test for successful amplification. Once amplification was verified, large scale PCR was performed using the following materials and solutions: 200 M dNTPs (Promega), 1 M of each primer, 1.5 mM MgCl2, 0.25 units Platinum Pfx DNA polymerase (Invitrogen), buffer (Idaho Technology), and 3 L template DNA (Wong et al, 2015).

Amplification was performed on a Rapid Cycler (Idaho technologies) in glass capillaries.

Thermocycling for the two mitochondrial genes were COI; 3 min at 94 C followed by 35 cycles of 94 C for 1s, 56 C for 1s and 68 C for 20s, ND4; COI; 3 min at 94 C followed by 35 cycles of

94 C for 1s, 50 C for 1s and 68 C for 20s.

5 PCR cleanup, labeling, and Sanger sequencing was performed by Genewiz (115

Corporate Boulevard South Plainfield, NJ 07080). Forward strands were sequenced for both COI and ND4, in addition to reverse strands for COI. Sequences were trimmed using Finch TV 1.5.0

(Geospiza Inc.). Alignment was performed and edited by eye using Mesquite v3.11 (Madison and Madison, 2011). Once aligned, codon sequences were verified using Mesquite database for

Invertebrate mitochondrial coding. No stop codons were present in any individuals for either

COI or ND4.

Phylogenetic Analysis

The phylogenetic analysis was run using 111 individuals, including four individuals that are not identified as Spirobolomyia or Blaesoxipha. These four outgroup individuals, one individual each of Ravina anxia, Ravina derelicta, Fletcherimyia fletcheri and Mecynocorpus salvum, were selected as outgroups based on Stamper et al. (2013). 76 individuals were identified as Blaesoxipha, with 13 identified to species representing six subgenera. 31 individuals were identified as Spirobolomyia, of which six were identified as one of two species.

The relationship of Blaesoxipha and Spirobolomyia were inferred using Bayesian

Inference and Maximum Likelihood. Bayesian Inference was inferred using MrBayes v3.2.1

(Ronquist and Huelsenbeck 2003). Maximum Likelihood was inferred using RAxML v8.2.10

(Stamatakis 2014). Both genes and codon positions can have different rates of mutation.

Assumption of heterogeneity across codon position or between genes can lead to biases in phylogenetic analysis (DeBry 1999). To avoid these biases, the data set was partitioned and a gamma distribution was used to model within-partition rate heterogeneity. The gamma distribution model was selected based on the previous study by Wong 2015.

6 For MrBayes, sequences were partitioned into first, second and third codon positions with both genes combined for a total of 3 subsets. Partitions were determined based on previous studies using mtDNA genes (Wong 2015; Stamper 2008). The first positon was set to number of substitution types (nst) = 6. For the second and third positions sites nst = 2. First and third position sites modeled among-site variation using the gamma distribution model. The second positon site modeled among-site variation by estimating the proportion of invariable sites. MrBayes was ran for 1x107 generations and sampled every 1000 generations. The first

40% of samples were discarded as “burn-in.” Posterior probabilities were used for nodal support. All other priors and parameters where set to the programs default settings.

For RAXmL, sequences were partitioned in to first, second and third codon positions for each gene, for a total of 6 subsets. RAXmL was ran using the GTR-gamma distribution model (- m = GTRGAMMA) across all partitions. A rapid bootstrapping analysis (-f = a) was used with the random seed set to 12000 (-p = 12000). RAXmL was run using bootstrap resampling to assess nodal support using 1000 pseudoreplicates (-# = 1000). For both MrBayes and RAXmL, only R. anxia (AT45) was named as outgroup.

Results

DNA sequence data was obtained for 108 individuals; 76 Blaesoxipha and 31

Spirobolomyia. The 76 individuals of Blaesoxipha included representatives of six of the ten currently recognized subgenera, all of which have a New World distribution. This data includes no representatives for those subgenera with only an Old World or Antillean distribution. The final alignment consisted of 1693 base pairs; COI was 1160 base pairs and ND4 was 533 base

7 pairs. The COI sequence had total of 459 variable base pairs; 91 in the first codon positon, 35 in the second codon position and 333 in the third codon position. The ND4 sequence had a total of 221 variable base pairs; 48 in the first codon position, 25 in the second codon position and

148 in the third codon position.

Bayesian Inference and Maximum Likelihood inferred very similar relationships among all individuals. Both trees found Spirobolomyia to be monophyletic (Fig. 1; BPML=77; PP=1.0).

The Bayesian Inference and Maximum Likelihood different in their placement of the unidentified individual Blaesoxipha sp. (AW49). The Bayesian tree supported Blaesoxipha sp.

(AW49) as a sister clade to all other Blaesoxipha (PP=0.95). In the Maximum Likelihood analysis,

Blaesoxipha sp. (AW49) was nested within the other Blaesoxipha clades. However, the nodes in the Maximum Likelihood were unsupported. Both trees also show a clade of unidentified

Spirobolomyia (Fig. 1; BPML=98; PP=1.0).

The results of the analysis returned an unsupported monophyletic Blaesoxipha in the

Maximum Likelihood analysis, however the monophyly was well supported in the Bayesian analysis. Within Blaesoxipha, some of the named subgenera are supported as being monophyletic while others are not. The subgenus Acridiophaga was paraphyletic. Within the clade containing Acridiophaga, the representatives from two additional subgenera, Tephromyia and Servaisia, were present. While Blaesoxipha augustifrons and Blaesoxipha taediosa are both apart of the subgenus Acridiophaga, the analysis supports Blaesoxipha

(Acridiophaga)augustifrons being more closely related to Blaesoxipha (Servaisia) uncata (Fig 1;

BPML=100; PP=1.0) and both being distantly related to Blaesoxipha (Acridiophaga) taediosa with high nodal support from both analyses (Fig. 1; BPML=80; PP=1.0).

8 At the species level, all the nodes for Spirobolomyia were well supported by both analyses. The clade of Blaesoxipha plinthopyga was well supported as monophyletic. The monophyly of the clade encompassing Blaesoxipha cessator has high Bayesian support, but low support by the Maximum Likelihood analysis. Blaesoxipha mex was monophyletic, however there is no nodal support from either the Bayesian inference or Maximum Likelihood. The monophyly of Blaesoxipha caridei, is not supported, though there is strong support for several clades within the species.

With exception of the 14 unidentified Spirobolomyia individuals that formed a clade with strong support, a majority of the other unidentified individual fell within clades of individuals with morphological identifications. There were only eight unknown individuals that did not fall within a clade of known individuals

9 Spirobolomyia

Blaesoxipha

Fig 1. The inferred Bayesian phylogeny of Blaesoxipha and Spirobolomyia using mtDNA genes COI and ND4. Support values for nodes are given in the following order: bootstrap values from ML analysis using RAXmL/Bayesian posterior probabilities using MrBayes. Subgenera are indicated by color. All bootstrap values under 50 and posterior probabilities under 0.95 are indicated by a (-). No nodal support values were indicated for nodes that were unsupported by both analyses.

10

Discussion

Spirobolomyia was monophyletic. However, due to the contradicting support from the two analyses for the monophyly of Blaesoxipha, it is difficult to rule out Spirobolomyia as a subgenus of Blaesoxipha. There are 242 species and 10 subgenera of Blaesoxipha for which this study only contained 13 different known species across six subgenera. The only subgenus with a

New World distribution not represented in the study was subgenus Kellymyia. Subgenera

Aldrichisca, Abapa and Blaesoxipha have distribution of either Old World or are restricted to the Antilles. Since the subgenera not in the study do not have the same distribution as

Spirobolomyia, I expect that adding these subgenera would offer little insight into the relationship of Spirobolomyia and Blaesoxipha.

The monophyletic clade of Spirobolomyia is comprised of six individuals morphologically identified as Spirobolomyia flavipalpis, two as Spirobolomyia singularis and 24 unidentified individuals. Spirobolomyia flavipalpis and 10 of the unidentified individuals form a monophyletic clade. The two single Spirobolomyia singularis form a monophyletic clade. The monophyletic clade of unidentified Spirobolomyia suggest the presence of a third species in this study (Hennig 1966). Of the 14 individuals in this subclade, none have been morphologically identified. I suspect it these individuals are one of the three Spirobolomyia other than

Spirobolomyia flavipalpis and Spirobolomyia singularis. Further investigation into these fourteen individuals and a morphological identification will need to be made to determine which species of Spirobolomyia this clade may represent.

11 Blaesoxipha was supported as a monophyletic clade by Bayesian Inference but had no support by Maximum Likelihood. Individual AW49 showed as a sister clade to all Blaesoxipha in the Bayesian tree but was nested within Blaesoxipha on the Maximum Likelihood tree. I reexamined the sequences for individual AW49. While the sequence quality was high both COI

(861 bp) and ND4 (632 bp) were short sequences. When the sequences were run through

BLAST (Altschul et al.), both sequences returned at 93% match to individual Blaesoxipha cessator (AW27). I suspect this disagreement in the phylogenies would be resolved if I had longer sequences for AW49 or even removed AW49 from the analyses.

The subgenus Gigantotheca formed a monophyletic clade with several subclades. All individuals of Blaesoxipha plinthopyga, including seven unidentified individuals, formed a monophyletic clade with strong nodal support. Blaesoxipha cessator, along with eight unidentified individuals, formed a monophyletic clade with strong Bayesian Inference support.

The unidentified individuals that formed a monophyletic clade with those that were morphologically identified will lend insight into the possible identification of those unknown individuals.

Subgenus Acanthodotheca has strong nodal support for its monophyly. Blaesoxipha alcedo and Blaesoxipha eleodis both form monophyletic clades. However, Blaesoxipha arizona is paraphyletic, with a nested clade of Blaesoxipha masculina and three individuals forming a single clade. Upon further investigation of the identification by Greg Dahlem, it was concluded that morphologically these two are very similar making it extremely difficult to identify the two- separate species.

12 The subgenus of Acridiophaga was found to be paraphyletic with strong support on several nodes. It was inferred that Blaesoxipha (Acridiophaga) augustifrons is more closely related to Blaesoxipha (Servaisia) uncata and both are equally related to Blaesoxipha

(Acridiophaga) taediosa. This clade also includes Blaesoxipha (Tephromyia) hunteri. There is also a lack of monophyly within the Blaesoxipha caridei clade. With the high nodal support of the subclades, there is a large possibility the Blaesoxipha caridei clade may contain more than one species.

Conclusion

The purpose of this study was to determine whether mtDNA would support a two-genus classification or suggest reducing Spirobolomyia to a subgenus of Blaesoxipha. While

Spirobolomyia is monophyletic, contradicting support for the monophyly of Blaesoxipha make me unable to draw a conclusion about the generic status of Spirobolomyia. More information such as, more identified individuals, different mtDNA genes and genomic DNA are needed to infer the relationship of Spirobolomyia and Blaesoxipha.

13 Table 1. Specimen Identification: Genus, subgenus, species and voucher

Genus Subgenus Species Voucher

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) BG46

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) AE50

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) AK02

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) AJ80

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) AJ81

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) BG49

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) AH77

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) BG66

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) AG29

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) AC79

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) BG50

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) BG47

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) BH76

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) BG48

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) BH77

Blaesoxipha Gigantotheca plinthopyga (Wiedemann) F1

Blaesoxipha Acridiophaga caridei (Brethes) AW28

Blaesoxipha Acridiophaga caridei (Brethes) AW33

Blaesoxipha Acridiophaga caridei (Brethes) AW44

14 Blaesoxipha Acridiophaga caridei (Brethes) AX65

Blaesoxipha Acridiophaga caridei (Brethes) AX68

Blaesoxipha Acridiophaga caridei (Brethes) AY07

Blaesoxipha Acridiophaga caridei (Brethes) AY10

Blaesoxipha Acridiophaga caridei (Brethes) AW29

Blaesoxipha Acanthodotheca eleodis (Aldrich) AW37

Blaesoxipha Acanthodotheca eleodis (Aldrich) AY04

Blaesoxipha Acanthodotheca alcedo (Aldrich) AY08

Blaesoxipha Acanthodotheca alcedo (Aldrich) AY09

Blaesoxipha Acanthodotheca alcedo (Aldrich) AY25

Blaesoxipha Gigantotheca mex (Pape) AX79

Blaesoxipha Gigantotheca mex (Pape) AX81

Blaesoxipha Gigantotheca mex (Pape) AY23

Blaesoxipha Gigantotheca cessator (Aldrich) BI21

Blaesoxipha Gigantotheca cessator (Aldrich) BI25

Blaesoxipha Gigantotheca cessator (Aldrich) AW27

Blaesoxipha Gigantotheca cessator (Aldrich) AW38

Blaesoxipha Acanthodotheca masculina (Aldrich) AX74

Blaesoxipha Acanthodotheca masculina (Aldrich) AW35

Blaesoxipha Acridiophaga taediosa (Aldrich) AY24

Blaesoxipha Acridiophaga augustifrons (Aldrich) AY01

15 Blaesoxipha Servaisia uncata (Wulp) AS39

Blaesoxipha Acanthodotheca arizona (Pape) AW31

Blaesoxipha Acanthodotheca arizona (Pape) AW32

Blaesoxipha Acanthodotheca arizona (Pape) AW36

Blaesoxipha Acanthodotheca arizona (Pape) AX66

Blaesoxipha Acanthodotheca arizona (Pape) AX73

Blaesoxipha Acanthodotheca arizona (Pape) AY11

Blaesoxipha Speciosia speciosa (Lopes) AW71

Blaesoxipha Tephromyia Hunteri (Hough) AW40

Blaesoxipha sp. AD05

Blaesoxipha sp. AW49

Blaesoxipha sp. AW51

Blaesoxipha sp. AW52

Blaesoxipha sp. AW53

Blaesoxipha sp. AW54

Blaesoxipha sp. AW56

Blaesoxipha sp. AW58

Blaesoxipha sp. AY26

Blaesoxipha sp. AY28

Blaesoxipha sp. AY29

Blaesoxipha sp. AY30

16 Blaesoxipha sp. AY31

Blaesoxipha sp. AY32

Blaesoxipha sp. AY33

Blaesoxipha sp. AZ02

Blaesoxipha sp. AZ03

Blaesoxipha sp. AZ04

Blaesoxipha sp. AE30

Blaesoxipha sp. AE53

Blaesoxipha sp. AG41

Blaesoxipha sp. AG56

Blaesoxipha sp. AJ11

Blaesoxipha sp. AJ24

Blaesoxipha sp. AK10

Blaesoxipha sp. BI76

Blaesoxipha sp. BI68

Spirobolomyia sp. BK33

Spirobolomyia sp. BL31

Spirobolomyia singularis (Aldrich) AA47

Spirobolomyia singularis (Aldrich) AB06

Spirobolomyia flavipalpis (Aldrich) AV20

Spirobolomyia flavipalpis (Aldrich) AV21

17 Spirobolomyia flavipalpis (Aldrich) BK26

Spirobolomyia flavipalpis (Aldrich) BK27

Spirobolomyia flavipalpis (Aldrich) BL42

Spirobolomyia flavipalpis (Aldrich) BL76

Spirobolomyia sp. AB08

Spirobolomyia sp. AB09

Spirobolomyia sp. AB11

Spirobolomyia sp. AB12

Spirobolomyia sp. AB13

Spirobolomyia sp. AB14

Spirobolomyia sp. AB16

Spirobolomyia sp. AB18

Spirobolomyia sp. AB19

Spirobolomyia sp. AB20

Spirobolomyia sp. AB22

Spirobolomyia sp. AB23

Spirobolomyia sp. AB24

Spirobolomyia sp. AB27

Spirobolomyia sp. AB28

Spirobolomyia sp. AB29

Spirobolomyia sp. AB31

18 Spirobolomyia sp. AB32

Spirobolomyia sp. AB33

Spirobolomyia sp. AB34

Spirobolomyia sp. AB43

Spirobolomyia sp. AL77

Table 2. List of primers used for this study Primer Fragment Sequence Citation TY-J-1460 COI tacaatttatcgcctaaacttcagcc Simon et al., 1994 C1-N-2191 COI cccggtaaaattaaaatafaaacttc Simon et al., 1994 CI-J-1751F COI ggagcyccwgatatagchttc Debry et al., 2012 L2-N-3014 COI taatatggcagattactccattgga Stamper et al., 2013 N4-J-8502 ND4 gttggaggagctgctatattag Simon et al., 1994 N4-N-9194 ND4 attttttgaaagaagtttaattcc Yu et al., 1999

19 References

Aldrich, J. M. (1916). Sarcophaga and allies in North America, Entomological Society of America.

(Murphy-Divins Co. Press: Lafayette, IN, USA.)

Altschul, S.F., Gish, W., Miller, W., Myers, E.W. & Lipman, D.J. (1990). "Basic local alignment search

tool." J. Mol. Biol. 215:403-410.

Allen, G., Pape, T. (1996). Description of Female and Biology of Blaesoxipha ragg Pape (Diptera:

Sarcophagidae), a Parasitoid of Sciarasaga quadrata Rentz (Orthoptera: Tettigoniidae) in

Western Australia. Australian Journal of Entomology

1996 vol: 35 (2),147-151

Crouzel, I. S., Salavin, R. G. (1961). Contribucion a la biologia de Sarcophagidae (Insecta, Diperta) Rev.

Inv. Argic. Buenos Aires 15,649-658

DeBry, R. W. (1999). Maximum likelihood analysis of gene-based and structure-based process

partitions, using mammalian mitochondrial genomes. Systematic biology, 48(2), 286-299.

DeBry, R., Timm, A., Wong, E., Cookman, C., Stamper, T. & Dahlem, G. A. (2012). DNA-Based

Identification of Forensically Important Lucilia (Diptera: Calliphoridae) in the Continental United

States*. Journal of Forensic Science, doi: 10. 1111/j. 1556-4029. 2012. 02176. x

Downes, W.L. Jr., (1965). Family Sarcophagidae-Pp. 933-961 in Stone, A. et al., eds. A catalog of Diptera

of America north of Mexico. United Stated Dept Agric., Agricultural Handbook 276, iv + 1696 pp.

Washington

Fitch, W. M., Margoliash, E. (1967). A method for estimating the number of invariant amino acid coding

positions in a gene using cytochrome C as a model case. Biochem Genet 1, 65–71.

20 Greathead, D. J. (1963). A review of the insect enemies of Acridoidea (Orthoptera). Trans. R. Ent. SOC.

Lond. 114, 437-517.

Hennig, W. (1966). Phylogenetic Systematics. Univ. Il- linois Press, Urbana, Illinois.

Hillis, D. M., Bull, J. J. (1993). An empirical test of bootstrapping as a method for assessing confidence

in phylogenetic analysis. Systematic biology, 42(2), 182-192.

Kelly, E. G. (1914). A new sarcophagid parasite of grasshoppers. J. agric. Res. 2: 435-446

Leonide, J. (1964). Contribution a I’etude biologique des dipteres sarcophagides parasites d’acridiens.

Ponte de larves et infestation de l’hbte par le Blaesoxipha berolinensis Vill. C. R. Acad. Sc. Paris

258, 4352-4354.

Leonide, J. (1967). Contribution a l’etude biologique des diptkres sarcophagides parasites d’acridiens.

111: Cycle biologique de Blaesoxipha rossica Vill., injection de larves dans le corm de I’hbte uar

les femelles de sarcophaaides. Max Netherv nenerouslv Drovided C.R. Acad: sc. Paris i65, 232-

234.

Leonide, J., Leonide, J.C. (1986). Les dipteres sarcophagidks endoparasites des orthopteres franqais.

Essai &rase space and accommodation at Cape Naturilkte biotaxonomique. Universite de

Provence.

Lopes, H. S., (1975). On some North American Sarcophagidae with red legs (Diptera). Revista brasileira

de biologia., 35 (1), 155-164

Maddison, W.P., Maddison, D.R. (2011). ‘Mesquite 2.73: a modular system for evolutionary analysis.’

Available from [http://mesquiteproject.org].

21 Middlekauf, W. W. (1958). Biology and ecology of several species of California rangeland grasshoppers

(Orthoptera: Acrididae). Pan-Pacif. Ent., 34, 1-11

Pape, T. (1990). Revisionary notes on American (Diptera: Sarcophagidae). Tijdschrift

voor Entomologie, 133(1), 43-74

Pape, T. (1994). The world Blaesoxipha Loew, 1861 (Diptera: Sarcophagidae). Ent. scand. Suppl. 45, 1-

247.

Pape, T. (1996). Catalogue of the Sarcophagidae of the World (Insecta: Diptera), Memoirs on

Entomology International, Vol. 8. Associated Publishers, Gainesville, FL.

Roback, S. S. (1954). The Evolution and of the Sarcophaginae (Vol. XXIII). Urbana: The

University of Illinois Press.

Ronquist, F., and Huelsenbeck, J. P. (2003). MrBayes 3: Bayesian phylogenetic inference under mixed

models. Bioinformatics, 19, 1572-1574.

Salvain, R. J. (1958). Notas biologicas sobre la mosca Servaisia (Protodexia) arteagai

(Blanch.) Rob. (Diptera, Sarcophagidae) parasito de la tucura. Revta Invest. Agric., 12, 299-309

Simon, C., Frati, F., Beckenbach, A., Crespi, B., Liu, H. & Flook, P. (1994). Evolution, weighting, and

phylogenetic utility of mitochondrial gene sequences and a compilation of conserved

polymerase chain reaction primers. Annals of the Entomological Society of America, 87, 651-

701.

22 Stamatakis, A. (2014). RAxML version 8: a tool for phylogenetic analysis and post-analysis of large

phylogenies. Bioinformatics, 30(9), 1312-1313.

Stamper, T. (2008). Improving the Accuracy of Postmortem Interval Estimations Using Carrion Flies

(Diptera: Sarcophagidae, Calliphoridae and Muscidae). (Electronic Thesis or Dissertation).

Retrieved from https://etd.ohiolink.edu/

Stamper, T., Dahlem, G. A., Cookman, C., & Debry, R. W. (2013). Phylogenetic relationships of flesh flies

in the subfamily Sarcophaginae based on three mtDNA fragments (Diptera:

Sarcophagidae). Systematic Entomology, 38(1) ,35-44.

Tourasse, N.J., Gouy, M. (1997). Evolutionary distances between nucleotide sequences based on the

distribution of substitution rates among sites as estimated by parsimony. Mol Biol Evol 14,287–

298.

Wong, E. (2015). DNA-based Species Delimitation of the Agriculturally Important Genus,

(Diptera: Sarcophagidae). (Electronic Thesis or Dissertation). Retrieved from

https://etd.ohiolink.edu/

Yu, H., Wang, W., Fang, S., Zhang, Y. P., Lin, F. J., Geng, Z. C. (1999) Phylogeny and evolution of the

Drosophila nasuta subgroup based on mitochondrial ND4 and ND4L gene sequences. Molecular

Phylogenetics and Evolution, 13, 556-565.

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