The Status of the Kisatchie Painted Crayfish (Faxonius Maletae) in Louisiana

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Spring 3-22-2019 The tS atus of the Kisatchie Painted Crayfish (Faxonius maletae) in Louisiana Jade L.M. McCarley

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THE STATUS OF THE KISATCHIE PAINTED CRAYFISH (FAXONIUS MALETAE) IN LOUISIANA

JADE L. M. MCCARLEY

A thesis submitted in the partial fulfillment of the requirements for the degree of Master of Biology Department of Biology

Lance R. Williams, Ph.D., Committee Chair

College of Arts and Sciences

The University of Texas at Tyler May 2019

ACKNOWLEDGMENTS

I would like to acknowledge Dr. Bob Wagner, Quantitative Ecological Services

(QES), Jody Patterson, Sarah Pearce, and the staff at Fort Polk for all of their support throughout this project. I am sincerely grateful for my advisor, Lance Williams for this opportunity. He has guided me through graduate school and has inspired me to continue as a scientist. Thank you, Marsha Williams for your dedication to field sampling and especially for welcoming me into your family while at UT. Thank you, Josh Banta, for lending your genetic guidance, educating me on Rad-Seq, and for being patient with me throughout the project. Thank you, John Placyk for your assistance with DNA extraction processes and phylogenetic analyses. Thank you, Kate Hertweck for all of the coding assistance and guidance during data analysis. Thank you to Brent Bill for providing guidance when called upon. From the bottom of my heart, I would like to extend the utmost thanks to my technicians Tyler Grabski and Ivy Sudduth for enduring the elements and always being reliable company during sampling and lab work. I would also like to extend my gratitude to Rachel Quintanilla and Katy Lunceford for their assistance in the field. Thanks to fellow graduate students Matt Wolkov, Bridget Fitzgerald, and

Nevada King for supporting me during the rigor of graduate school. I would like to personally thank my mother Shawn McCarley, brother Steven Holland, and fiancé Kyle

Mason for their constant support during my research conquest while away from home.

Without you three, I would not possess the drive and dedication I needed to complete my

Master’s Degree in Biology. A large amount of team work made this research possible. I will forever be grateful and thankful for this experience.

TABLE OF CONTENTS

List of Figures…………………………...... ……………………………………………...iii

List of Tables.…………………………………………………………………………….iv

Abstract………………………………………………………………...…………………iii

Chapter One: Introduction….…………………………………………..…………………1

Classification……………………………...……………………………………….2

Reproduction and Mating Behavior……………………………………………….2

Historical Presence in Texas………………………………………………………3

Historical Presence in Louisiana…………………………………………………..4

Chapter Two: Materials and Methods…………………………………...………………..6

Study Area………………………………………………………………………...6

Field Survey Methods……………………………………………………………11

Species Morphology and Identification…………………………………….……13

Population Dynamics……………………………………………………….……14

DNA Extraction and Quantification……………………………………………..15

Restriction Site Associated DNA Sequencing……………………………..….…16

Phylogenetic Analyses (Maximum Likelihood Approach)……………………...17

Phylogenetic Analyses (Bayesian Clustering Approach)………………………..19

Phylogenetic Analyses (AMOVA)………………………………………………20

Chapter Three: Results…………………………………………………...………………21

Population Dynamics…………………………………………………………….21

Restriction Site Associated DNA Sequencing……………………………….…..23

Phylogenetic Analyses (Maximum Likelihood Approach)…...…………………24

Phylogenetic Analyses (Bayesian Clustering Approach)………………………..26

Phylogenetic Analyses (AMOVA)…………………………………………...….27

Chapter Four: Discussion……….……………………………………….……………….28

References………………………………………………………………………….…….33

Appendix A………………………………………………………………………………42

LIST OF FIGURES Figure 1. Historical sampling locations for the Kisatchie Painted Crayfish, Faxonius maletae, in Texas………………………………………………………………...………..7

Figure 2. Texas and Louisiana historical sites sampled for the Kisatchie Painted Crayfish

(Faxonius maletae)………………………………………………………………………..8

Figure 3. Map of Fort Polk near Leesville, Louisiana that indicates the boundaries of both

Fort Polk and the Peason Ridge Training Area (PRTA)………………………………….9

Figure 4. Map showing the tributaries found within the Peason Ridge Training Area

(PRTA) located in the Red River Drainage basin of Louisiana...... ………………...... 10

Figure 5. Pictures of Kisatchie Painted Crayfish, Faxonius maletae, indicating coloration patterns…………………………………………………………………………………...14

Figure 6. Map of sites where Kisatchie Painted Crayfish, Faxonius maletae, were collected in Texas and Louisiana for this project….…………………………...………..21

Figure 7. Length (mm)-weight (g) relationship for Kisatchie Painted Crayfish, Faxonius maletae……………………………………………………………...……………………22

Figure 8. Size class distribution for Kisatchie Painted Crayfish, Faxonius maletae….…23

Figure 9. Maximum likelihood phylogeny with proportional branch lengths for Faxonius maletae...…………………………………………………………………………………25

Figure 10. STRUCTURE plot results indicating inferred ancestry between Texas and

Louisiana subpopulations of Faxonius maletae (K=1)…………………………………..26

iii

LIST OF TABLES

Table 1 AMOVA results among populations, individuals within populations, and within individuals for Texas and Louisiana subpopulations of Faxonius maletae (average over

41 loci)……………………………………………….…………………………………..27

Table 2 Texas historical sites for Faxonius maletae sampled in 2017—2018…………..42

Table 3 Louisiana historical sites for Faxonius maletae sampled in 2017—2018………43

Table 4 Sites sampled for Faxonius maletae on Fort Polk military base in Leesville, LA in 2017—2018………………………………………………………………...…………44

Table 5 Average Maximum Log Likelihood values provided from STRUCTURE for each

Faxonius maletae individual.…………………………………………………………….44

ABSTRACT

THE STATUS OF THE KISATCHIE PAINTED CRAYFISH (FAXONIUS MALETAE) IN LOUISIANA Jade L. M. McCarley

Thesis Chair: Lance R. Williams, Ph.D.

The University of Texas at Tyler May 2019

The Kisatchie Painted Crayfish, Faxonius maletae, are considered to be imperiled and potentially endangered in Texas and Louisiana. There are two known subpopulations, and previous work suggested these subpopulations may be highly genetically differentiated and therefore deserving of different subspecies or species status. Upon field sampling and performing DNA extractions, Restricted Site Associated

DNA Sequencing (RAD-Seq) was performed based on Single Nucleotide Polymorphisms

(SNPs) to assess genetic variability between Texas and Louisiana subpopulations.

Phylogenetic analysis indicated that the two subpopulations are not genetically differentiated from one another. Population genetic analyses further supported that the species are not genetically differentiated from one another (P=0.49, FST = 0.10). The results presented here suggest that the two subpopulations are still exchanging alleles with one another and are not separate species or subspecies. Faxonius maletae is declining indicating the importance for conservation status in Texas and Louisiana.

CHAPTER ONE: INTRODUCTION

Crayfish serve as keystone species in aquatic habitats and conservation efforts should focus on them as indicators for habitat quality (Crandall 2007). Approximately

48% of freshwater crayfish species are considered vulnerable, threatened, or endangered

(Taylor et al. 2007). The Kisatchie Painted Crayfish (Faxonius maletae) are primarily stream dwellers (Penn 1952). They have been collected in small to moderately sized streams with bottoms composed of white sand or gravel that exhibit clear water. They have also been documented in large rivers with bottoms that are composed of mud that have extremely silty water (Walls 1985). Habitat characteristics such as water depth have been shown to influence the length of F. maletae. Those that occur in deep water have been documented to reach 101.6 mm and those that occur in shallow water rarely reach lengths over 50.8 mm long (Walls 2005). Both the Texas and Louisiana subpopulations occupy areas that have these habitat characteristics, however Texas habitats were more stagnant, muddy, and foul-scented than those in Louisiana.

Faxonius maletae is listed as a species of special concern by the Louisiana

Natural Heritage Program. Faxonius maletae was also included in a 2010 petition for listing of 404 species under the Endangered Species Act. It is listed by the International

Union for the Conservation of Nature (IUCN) as data deficient, and it is exhibiting a decreasing population trend (Adams et al. 2010). It has a Global Heritage Ranking of G2, or imperiled, in the Kisatchie Bayou of Louisiana because of population declines (Taylor et al. 2007); however, since there are little or no data on the ecology or population genetics of this species, its listing under the Endangered Species Act has been delayed.

Faxonius maletae has been an elusive species to study and little is known about their behavior, reproduction, diet, and general physiology. Little is known about conservation threats, habitat requirements, and the distribution of most North American crayfish species. Threats such as limited natural range, nonindigenous crayfish invasion, and habitat alteration are responsible for imperilment of crayfishes (Taylor et al. 2007).

During the winter season, crayfish exhibit a higher mortality as mating stress, starvation, and predation rates increase while environmental effects such as low dissolved oxygen levels and temperature fluctuations can also influence mortality (Kichler 1984). Prior crayfish studies indicate that females become more resistant to capture the colder it gets, reiterating winter sampling difficulty (Somers and Stechey 1986). This research project set out to provide data that may help in conserving the species if warranted.

Classification— The Painted Crayfish, Orconectes difficilis, (Faxon 1898) comprises four subspecies in the Orconectes genus. Orconectes difficilis in southeastern

Oklahoma, the Kisatchie Painted Crayfish, O. difficilis maletae, in areas of east Texas and west Louisiana, Orconectes difficilis hathawayi (Penn 1952) in east-central

Louisiana, and the Calcasieu Crayfish, O. blacki, in southwestern Louisiana (Walls

1985). Orconectes d. maletae was later renamed O. maletae in 1972 (Walls 1972) and was reclassified to Faxonius maletae in 2017 (Crandall and Fitzpatrick 1996; Fetzner

1996; Crandall et al. 2000; and Taylor and Knouft 2006; Crandall and De Grave 2017).

Reproduction and Mating Behavior—Most North American crayfish are documented to mate in early spring, but F. maletae is hypothesized to have a breeding peak in September and October (Walls 1985). This timeframe is when males are sexually mature and their 1st pleopods are horny, brown, and have sharp tips. This stage is

2 referred to as Form I. Form I males are visibly active in full view upon debris lining the stream while actively “displaying” their yellow chelae fingers. Females have not been documented to display chelae indicating that males display to attract female mates (Walls

1985). By the end of October, sperm plugs were present in females, indicating that mating had occurred recently (Walls 1985). Sperm plugs are deposited into female spermatheca and work to inhibit other males from mating with the female again (Pehnke and Pyae 2012). Females generally secrete a substance called glair before they release eggs. This substance allows the sperm plug to be dissolved at which point sperm from the spermatheca is distributed into the glair. The eggs pass through the glair for external fertilization and then attach to the pleopods of the females until they hatch and are developed enough to swim on their own (Clifford 1991).

Eggs hatch around May as juveniles tend to be most abundant in June and July.

Form II males (non-reproductive) exhibit 1st pleopods that are flabby, white, and rounded

(Walls 1985). Juvenile males molt to Form I and stay in this form through winter. In

March, Form I males molt to Form II to become non-reproductive until the coming fall when they will molt again, back to Form I (Walls 1985). The largest males tend to be found in the summer and it is speculated that males die after two years, in their second winter (Walls 1985). From prior research, the smallest observed Form I male had a total length, from the rostrum to the end of the telson, of 14 mm. The maximum total length collected during the same study was 44 mm (Walls 1985).

Historical Presence in Texas— Historical collecting localities were obtained from the Texas Parks and Wildlife Department. Sampling methods included exhaustive methods such as minnow traps baited with hotdogs (Hobbs and Lodge 2010) and dip

3 netting along banks and in riffles (Rabeni 1997). Faxonius. maletae were found to be absent in 14 of 24 Texas historical sites sampled (Brown 2017). One of the 25 total historical sites were inaccessible which prevented it from being sampled (Brown 2017).

Faxonius maletae was determined to be absent in 60% of its historical range in Texas

(Williams et al. 2014). The absence of F. maletae at these sites was caused by declining population numbers or an inability of the species to be detected by evasion (Brown

2017).

Mitochondrial DNA analysis using Cytochrome C Oxidase Subunit I (COI), the

16S mitochondrial gene, and the nuclear gene segment GADPH has been used to determine the phylogenetic relationship of F. maletae subpopulations in Texas and

Louisiana (Mathews 2008; Brown 2017). A high rate of mutation is exhibited in mitochondrial DNA (White et al. 2008). As a result of maternal inheritance and the lack of recombination, mitochondrial DNA is useful in showing phylogenetic relationships and determining whether species are genetically differentiated (Munasinghe et al. 2003;

Crandall et al. 2007). The mitochondrial DNA analysis suggested that the Texas and

Louisiana subpopulations have undergone divergence and are potentially undergoing allopatric speciation (Brown 2017).

Historical Presence in Louisiana—The Louisiana subpopulation found in the

Kisatchie National Forest was examined in prior research in central-western Louisiana from 1966-1970 by Jerry Walls (Walls 1985). The specimens were captured utilizing a dipnet and were also captured by hand around areas littered with debris or logs (Walls

1985). In 2005, F. maletae was determined to be absent in 4 out of 10 historical sites, indicating a decline (Walls 2005).

The primary focus of this research was to determine whether the Louisiana and

Texas subpopulations differ genetically using full genome analysis rather than mitochondrial DNA analysis. The Texas and Louisiana historical populations of

Faxonius maletae are thought to be isolated from one another by geographic barriers that have segregated the Red River from the Atchafalaya River such as the Great Raft, the causative agent for the formation of Caddo Lake (Bagur 2001; Holbrook 2007). A geographic distance of approximately 150 km separates the two subpopulations in

Natchitoches Parish, Louisiana and those found in Franklin County, Texas.

CHAPTER TWO: MATERIALS AND METHODS

Study Area— There are only two known subpopulations of this species. The species has been observed in high numbers within the boundaries of Fort Polk, Louisiana prior to 2017. One subpopulation resides in eastern Texas within the Cypress Creek and

Caddo Lake drainages (Figure 1). The other subpopulation resides in western-central

Louisiana within the Kisatchie Bayou and Red River drainages. Historical sites within

Texas and Louisiana visited during this study are shown in (Figure 2).

Within Texas ten historic sites were sampled within the Cypress Creek and

Caddo Lake drainages. Habitat composition varied substantially per site. Some areas were heavily littered with construction rock, under highway overpasses or bridges that also had murky, stagnant water with little vegetation present. Other sites included bedrock bottomed creeks, with heavy vegetation along the banks, and a high flow rate with riffles that rushed around larger boulders. Several Texas sites were stagnant, swamp-like, and a foul scent accompanied the area. There were also larger riverine systems where water reached depths of 6-feet that had a high flow rate. The hypothesized preferred habitat of F. maletae includes areas with varying water depth between one and four feet, heavy leaf littered or cobble lined stream bottoms with logs and large rocks that assist ambush predation and cover tactics (Klecka and Boukal 2014).

This habitat composition provides a prime setup for ambush predation, as I hypothesize

F. maletae are ambush predators.

Figure 1. Historical sampling locations for the Kisatchie Painted Crayfish, Faxonius maletae, in Texas. Blue dots are sites where crayfish may be extirpated, and red dots are representative of specimens collected in 2014 (Brown 2017).

Figure 2. Texas and Louisiana historical sites sampled for the Kisatchie Painted

Crayfish (Faxonius maletae).

Upon review of prior research of these subpopulations, I revisited the 10 Texas sites that were found to be occupied by Brown (2017). I did not revisit sites that were determined to be unoccupied in 2017 as the last survey had been completed the same year that this research was to be continued (Brown 2017, unpublished data). The details for

8 sites I visited in Texas such as site identification labels, counties, GPS coordinates, specimen identification labels, and the dates sampled can be found in Appendix (Table

1). I revisited the 10 Louisiana historical sites where F. maletae had older records of existence or had been documented to inhabit in 2005 (Walls 2005). I also revisited another site in Louisiana determined to be occupied by F. maletae, for a total of 11

Louisiana sites visited (Lance Williams, pers. obs). I revisited these sites to further assess population declines. The details for sites I visited in Louisiana such as site identification labels, GPS coordinates, specimen identification labels, and the dates sampled can be found in Appendix (Table 2). Apart from sampling at historical sites, I also sampled streams located on the military base Fort Polk in Louisiana. There are approximately 51,000 acres of land in the heaviest impacted area within Fort Polk known as the Peason Ridge Training Area (PRTA) in Louisiana shown in Figure 3.

Figure 3. Map of Fort Polk near Leesville, Louisiana that indicates the boundaries of both Fort Polk and the Peason Ridge Training Area (PRTA) (Williams et al. 2014).

The Fort Polk sites were targeted to determine occupancy in four tributaries

(Little Sandy, Odom, Tiger, and Lyle’s Creeks). The four tributaries all flow into the

Kisatchie Bayou that is a part of the Red River drainage basin (Figure 4).

Figure 4. Map showing the tributaries found within the Peason Ridge Training Area

(PRTA) located in the Red River Drainage basin of Louisiana (Williams et al. 2005).

This Red River basin eventually flows into the Mississippi River. Faxonius maletae had been collected in these four sites off highway 117 located near Little Sandy

Creek during routine stream biomonitoring (Williams et al. 2014). The four tributaries sampled within Fort Polk were not prior documented historical sites but are in the same water system as other historical localities (Walls 2005). As a result of detecting this species in these four tributaries, those sites can now be documented as sites where F. maletae are present. Fort Polk, LA site identification labels, counties, GPS coordinates,

10 specimen identification labels, and the dates sampled can be found in Appendix (Table

3).

Field Survey Methods—The most appropriate sampling time for F. maletae is suggested to be from April to October and therefore was chosen as a result of literature review (Walls 1972; Walls 1985; Walls 2005; Brown 2017). This time frame was chosen to maximize our chances of capturing enough specimens to complete genetic analysis. I did not sample from November—March in efforts to mitigate negative impacts on the species that could result from sampling during a stressful time period (Nowicki et al.

2007). Prior sampling over the course of four years from December to January yielded no crayfish being captured and flooding over the course of February to May was documented to routinely prevent collection in the Louisiana sites (Walls 1985).

Sampling was not performed in September of 2017 as a result of heavy flooding and dangerous environmental conditions from Hurricane Harvey. Sampling was also not performed in July and August of 2018 as a result of Fort Polk training rotations that prevented access to the Fort Polk sites.

Collection methods for crayfish vary depending upon habitat type. Several methods were attempted such as baiting minnow traps, electrofishing, and using D-frame kick nets. Dipnets were chosen in prior studies and baiting was found to be inefficient

(Walls 2005). Electrofishing for crayfish has been considered the most efficient sampling method in the past that resulted in two to four times greater abundance estimates than using hand net methods (Rabeni et al. 1997). Electrofishing has also been shown to provide accurate species relative abundance estimations during analysis

(Meador et al. 2011).

A field crew consisting of students from the University of Texas at Tyler Biology

Department performed sampling for F. maletae utilizing all of the prior mentioned methods. At each Fort Polk site, a 100-meter transect was walked beginning at the culvert and we proceeded walking upstream away from the road to prevent disturbance.

A backpack electroshocker was utilized from August to October of 2017 and 2018. In

2017, we had an extremely low capture rate, potentially as a result of using the electroshocker. Voltage was adjusted when necessary due to low conductivity in these streams. A two-pass depletion method was utilized by using dip nets in combination with the electroshocker (Kimmel and Argent 2006). When utilizing a backpack electroshocker, F. maletae dashed away at a high speed and capture rates were low as a result. Crayfish tend to be nocturnal and this could have played a role in low capture rates as we sampled during the day. The water was also quite shallow in the areas sampled and

F. maletae inhabited areas cluttered with debris, logs, and deeply undercut banks. The water was also turbid, and we could not see the bottom of the streams. This made sampling with dipnets nearly impossible. Both of these methods were determined to be ineffective for our sampling sites. We utilized them again the next year during the same time frame as a second attempt to capture crayfish using these methods but were unsuccessful.

For the remainder of the sampling dates, minnow traps were baited with fried chicken tenders and tethered to debris lining undercut banks every 25 meters. This bait was chosen in hopes that the oil would dissipate through the water in attempt to attract more crayfish to the trap. The diffusion of the bait is dependent on water turbulence and bait type (Somers and Stechey 1986). There were 4 traps placed in each of the Fort Polk

12 sites. When we sampled the historic sites of Louisiana and Texas, we put up to 6 traps per stream to maximize capture rates and sometimes extended beyond the 100 meter transects if suitable habitat was seen nearby. Traps were placed in prime habitats that exhibited high capture rates over the course of the project. Most were placed in areas littered with heavy construction rocks and gravel bottoms with shallow, fast flowing water that included riffles. Not all sites had this type of habitat therefore some were placed in 4-feet-deep water with no bottom in sight and target crayfish were still captured there. These traps were left for 24 hours overnight. The traps were emptied after this time period and any F. maletae were removed for processing. For the sites visited, trapping was determined to be the most effective method for capturing the most male F. maletae.

Species Morphology and Identification—Each specimen was visually identified as F. maletae. The morphological characteristics exhibited by this species include an olive tinted carapace. The chelae have a gradient of coloration patterns beginning with red tips that fade into light blue, then fading to an olive base with black speckles covering the chelae. All joints of the walking legs and chelae are painted with red and there are red marks above the eyes on the carapace on this species. The cervical groove is a darker brown coloration and the carapace is olive in color. This olive color extends along the

13 carapace and the tail segments are bordered with darker stripes until reaching the telson, which is primarily dark brown and olive in color (Figure 5).

Figure 5. Pictures of Kisatchie Painted Crayfish, Faxonius maletae, indicating coloration patterns.

Population Dynamics—Males were sexed based on prior outlined identification such as examining the Form I or reproductive status of male gonopods as well as examining presence of pleopods or the “extra set” of swimmerets that are not present in females (Hobbs 1989). Females were sexed by glair, egg presence, or attached young and the lack of extra pleopods (Johnson 2010). The specimens were then weighed (g) and the total length (mm) from the tip of the rostrum to the end of the telson was recorded. These data were utilized in population dynamic estimates such as length- weight relationship and size class distribution for males and females.

DNA Extraction and Quantification—During field processing two walking legs were removed from each specimen for DNA sequencing. Crayfish can regenerate these appendages, ensuring that the declining species will not exhibit further mortality resulting from removal (Durand 1960). The samples were placed in vials of 95% ethanol. Each specimen was given a unique identification label based on capture location and sample number for that location. Two walking legs yielded enough DNA for the analysis for the majority of the samples collected.

Samples were stored at -20°F in a freezer for preservation until extraction. The samples were prepared using the GeneJet Genomic DNA Extraction protocol by

ThermoScientific. Once Proteinase-K was added to the samples, they were placed on a

ThermoScientific Thermal Rocker model 4637-1CE. Samples were maintained at 56ºC for a total of 24 hours. Two changes were made to the protocol. I increased the amount of time on the thermal rocker to ensure enough DNA would be removed from the keratin exoskeleton of the samples (Yanhe et al. 2011) and elution buffer was excluded from the extraction process and replaced with 200 µL of molecular grade water that had been heated to 56ºC in a hot water bath. The latter choice was made in efforts to prevent additional salt and contaminants from decreasing DNA purity.

After extraction the samples were quantified using a NanoDrop 2000 spectrophotometer (ThermoFisher Scientific) and a Qubit 4 Fluorometer (ThermoFisher

Scientific). Every extraction was tested using both pieces of equipment as the Qubit does not provide an estimation of DNA contaminants or salt levels, whereas the NanoDrop does (Desjardins et al. 2009). Using both pieces of equipment to read DNA samples

15 allows for the assessment of DNA purity and quantity of dsDNA of each sample

(Simbolo et al. 2013).

The NanoDrop was prepped by cleaning it with ethanol and Kim wipes.

Molecular grade water was used to zero the machine and then each sample was read by placing 5 µL of the sample on the NanoDrop. Each sample was assigned a total DNA concentration amount in ng/µL, A-260 10 mm path value, A-280 10 mm path value,

260/280 value, and 260/230 value by the NanoDrop. It was zeroed every ten samples to prevent error. The Qubit was also used to assign a DNA concentration amount for each sample. Each sample was 50 µL. A minimum of 20 ng/µL of DNA is required to be sequenced and samples must be normalized. Any sample that was well over this requirement had to be diluted using molecular grade water. Any sample that was under this requirement had to be concentrated using a Labconco Centrivap DNA Vacuum

Concentrator (LABCONCO). Samples were transferred to a 96-well plate. The well plate was packaged in a Styrofoam cooler with 4 pounds of dry ice and was shipped to

Floragenex for genomic data generation (Portland, OR).

Restriction Site Associated DNA Sequencing—The definition of phylogenetic analysis is the study of the evolutionary relationships of organisms. As phylogenetic studies focus on evolutionary relationships, they help with taxonomic classification

(Hedges 2002). Restriction Site Associated DNA Sequencing (RAD-Seq) is a Next

Generation Sequencing method that generates genome-wide polymorphic genetic data.

As single-gene phylogenies have been shown to be misleading when interpreting evolutionary relationships, assessing the SNPs from throughout the genome would better represent evolutionary relationships of Texas and Louisiana subpopulations of F. maletae

(Pamilo and Nei 1998; Gontcharov 2004; Maddison and Knowles 2006; Cariou 2013;

Spinks et al. 2013). The genomic DNA extracted from individual specimens are first introduced to a restriction nuclease that functions to digest the sample into DNA fragments that allow adapter attachment to said fragments. This process amplifies the sequences and applies tags for the process of high-throughput Illumina sequencing which identifies locations that contain Single Nucleotide Polymorphisms (SNPs) or a variation in a single base pair in a DNA sequence. Robust phylogenetic analysis can then be performed utilizing the SNP dataset to infer evolutionary relationships between individuals (Narum et al. 2013).

The SNP dataset provided by Floragenex was filtered to eliminate any monomorphic loci or DNA code that is the same for all individuals included in the analysis. The data set was further filtered to eliminate all but one locus per RAD-Seq fragment. This process eliminated redundant loci because neighboring loci frequently transmit the same alleles resulting in linkage disequilibrium among loci on the same fragments. The RAD-Seq analysis included 62 individuals, 49 individuals were those that I captured which included 8 females and 41 males; Six individuals from Texas and

43 individuals were from Louisiana. Ten individuals were outgroup samples that were provided by Neil Ford (UT Tyler). Eight individuals were male and 2 were female.

Three additional Faxonius maletae (Form I) male gonopod samples were processed and provided by Beau Gregory (Louisiana National Heritage Program). Floragenex provided the SNP dataset in an IUPAC file that was used in later analyses.

Phylogenetic Analyses (Maximum Likelihood Approach)— In this study two approaches for phylogenetic analysis were used. The first approach to assess

17 phylogenetic relationships between Texas and Louisiana populations of F. maletae was the Maximum Likelihood approach. This approach uses standard non-parametric bootstrapping to assess the accuracy of the estimated confidence level for the phylogenetic tree constructed (Efron et al. 1996). The confidence level assigned is calculated based on assigned maximum likelihood values. RAxML (Randomized

Axelerated Maximum Likelihood) is utilized to align SNPs and is frequently used to determine evolutionary relationships among individuals (Stamatakis 2014; Ogilvie et al.

2016).

Before using programs like RAxML an outgroup individual from the SNP dataset had to be chosen to use in the phylogenetic analysis. Phylogenetic studies utilize outgroups to assess evolutionary relationships. Most reconstruction methods produce phylogenetic trees that are unrooted meaning they cannot accurately infer relationships in relation to time. To prevent this issue, a tree must be rooted and an outgroup that is outside of the ingroup, or taxa under investigation, must be included in the analysis

(Philippe et al. 2011). For this study the Western Painted Crayfish, Orconectes palmeri longimanus, was utilized as an outgroup. I chose the outgroup sample OG10-R1-1 as my outgroup for this analysis as it had the most SNPs available in comparison to the other nine outgroup samples that were submitted for RAD-Seq analysis. The dataset used in

RAxML was filtered to remove all duplicate samples and all other outgroup SNP data.

This process removed 8 male and 1 female outgroup samples as well as 4 male and 2 female F. maletae.

RAxML uses the Evolutionary Placement Algorithm (EPA) and other reliable tree search algorithms to assign maximum likelihood values to the SNP dataset and place

18 short reads into a given reference phylogeny that was obtained from the full-length sequences. RAxML also uses an ascertainment bias corrections (ASC) when only variant

SNP sites are included in the alignment. This process returns trees that have been assigned reliable likelihood scores. These are used to determine the evolutionary origin of the reads (Berger et al. 2011; Stamatakis 2014). The RAxML analysis was performed over 47 individuals, 6 females and 40 male F. maletae, and 1 female outgroup.

Phylogenetic Analysis (Bayesian Clustering Approach)— The second approach used to assess phylogenetic relationships between Texas and Louisiana populations of F. maletae was the Bayesian Clustering approach. In the approach, relationships and gene flow between species can be determined by assessing the number of genetic clusters and how each individual’s membership relates to those clusters (Pritchard et al. 2000). The program assigns individuals to clusters by applying Markov Chain Monte Carlo (MCMC) estimation. This process randomly assigns individuals to a pre-determined number of groups and then variant frequencies are estimated in each group. Individuals are reassigned based upon these frequency estimations (Porras-Hurtado et al. 2013). This process is repeated for 10,000 times or the specified value entered by the user. For this project, the filtered SNP dataset was utilized, and all outgroups were removed including

OG10-R1-1 because the inclusion of an outgroup would only create an outlier in the data.

The burn-in and MCMC were set to 10,000 iterations.

The number of clusters (K) to be tested can be specified by the user where K also represents inferred ancestry (Blanco-Bercial and Bucklin 2016). For this study K was set from 1 to 14 and was performed with 3 iterations. The optimal K value was visualized and the K=1 model was chosen as it represented the lowest average log-likelihood value

19 over the 3 iterations for K=1 to K=14 (Rohlf and Sokal 1995). The average log likelihood values produced from STRUCTURE analysis can be found in the Appendix,

Table 4. The STRUCTURE analysis was performed over 46 individuals, 6 females and

40 male F. maletae.

Phylogenetic Analyses (AMOVA)—Like STRUCTURE, an Analysis of

Molecular Variance (AMOVA) is a genetic analysis that can provide information necessary to understand allele sharing patterns. AMOVA uses the amount of variance among groups (F-statistics) to determine whether there is a well-defined population structure. An AMOVA was performed over the filtered diploid data set, that had outgroups removed, using the program Arlequin Suite Version 3.5 (Excoffier and Lischer

2010). A locus by locus analysis with 1,000 permutations was completed. This process provided a fixation index (FST) value and P-Values for the Texas and Louisiana populations of F. maletae. F-statistics calculated by Arlequin are used to measure genetic differentiation among the species and the degree of inbreeding relative to a random mating population. F-statistics are also relative to the total genetic diversity of all samples (Wright 1965). The interpreted results from the AMOVA assesses the genetic similarity between two species. The FST reported includes the average of all pairwise comparisons of FST. The values range from 0 to 1 where zero indicates open gene flow meaning there is a higher amount of genetic diversity shared among populations investigated. The closer FST gets to one, the higher the degree of inbreeding. This would indicate decreased gene flow and genetic diversity between populations.

CHAPTER THREE: RESULTS

Population Dynamics—In Texas 10 historical sites were sampled and the

Kisatchie Painted Crayfish (Faxonius maletae) were only found in 2 out of 10 sites. In

Louisiana, 11 historical sites and 4 Fort Polk, LA sites were sampled and F. maletae were only found in 6 out of 11 sites (Figure 6).

Figure 6. Map of sites where Kisatchie Painted Crayfish, Faxonius maletae, were collected in Texas and Louisiana for this project.

For the Louisiana subpopulation length, weight, and sex data were utilized to make inferences about population dynamics. The length-weight relationship for F. maletae indicated that there is a linear relationship. The length in millimeters, regardless

21 of sex, is correlated with weight in grams. The R2 coefficient of determination is 0.87 indicating how well the fit of the data is to the line (p<0.001; Figure 7).

Figure 7. Length (mm)-weight (g) relationship for Kisatchie Painted Crayfish, Faxonius maletae.

The frequency of capturing males was much higher than that of capturing females

(X2 = 27.9, p<0.001; Figure 8). Based on a review of the data, this could be attributed to capture methods. All females were captured via a dip net or by using electrofishing in combination with a dipnet. All F. maletae that were captured in the minnow traps were males.

Figure 8. Size class distribution for Kisatchie Painted Crayfish, Faxonius maletae.

Restriction Site Associated DNA Sequencing—RAD-Seq analysis by Floragenex

(Portland, OR) yielded genome-wide data for the two population of F. maletae with total reads equaling 628,155,465. The mean reads per sample was 6,543,286.1. The number of quality-filtered RAD tags provided by the standard output of reads that passed FASTQ quality filters were 16,712,922, and the number of failing reads was 240,065. The total number of contigs extracted from the provisional clusters were 44,054, and the total number of contigs in the final assembly were 213,021 with an average base pair length of

92. The total cluster length was 19,597,932 bp.

Out of the 95 samples screened, the total number of candidate variants detected was 497,460, and the number of candidate variants filtered (because of missing or low-

23 quality data) was 496,863. The number of candidate variants passing all filters was 595.

The average number of polymorphisms within 200 bp of each variant was 5.2. The number of homozygous genotypes found was 39,518, and the number of heterozygous genotypes found was 13,242.

Phylogenetic Analyses (Maximum Likelihood Approach)—The phylogeny constructed based on SNP data, from Restriction Site Associated DNA Sequencing (Rad-

Seq), showed that the Texas and Louisiana subpopulations are intermixed and are not genetically differentiated (Figure 9). The rooted maximum likelihood tree indicates that the Texas and Louisiana populations are closely related. Bootstrap support for all nodes was low. The data was filtered to remove bootstrap support less than 70 for all individuals. The values are not observed on the tree nodes as a result of the filtering process.

Figure 9. Maximum likelihood phylogeny with proportional branch lengths for Kisatchie

Painted Crayfish, Faxonius maletae. Red: Louisiana, Blue: Texas, and Black: Outgroup.

A 0.005 scale bar is used to represent evolutionary lineages over time and is applied to branch lengths for determining relationships between individuals.

Phylogenetic Analyses (Bayesian Clustering Approach)— For the Bayesian cluster analysis of the Louisiana and Texas populations of F. maletae the most parsimonious number of inferred ancestral groups (K) was one. The Texas and Louisiana populations of F. maletae do not cluster separately from one another. The

STRUCTURE analysis indicated that inferred ancestry is 100% between the two subpopulations and they are not genetically differentiated, sharing a common ancestor.

The model chosen to represent the two subpopulations out of all models ranging from

K=1 to K=14 was the K=1 model. This model was chosen based on the lowest average log likelihood value between models over three iteration (Figure 10).

Figure 10. STRUCTURE plot results indicating inferred ancestry between Texas and

Louisiana subpopulations of Faxonius maletae (K=1).

Phylogenetic Analyses (AMOVA)—The AMOVA provided the percentage of molecular variation of Texas and Louisiana populations of F. maletae. These are explained by variation 1) among populations, 2) among individuals within populations, and 3) within individuals as represented in (Figure 11). The FST value of 0.10 represents the proportion of molecular variation among the Texas and Louisiana populations. This value indicates that there is no genetic differentiation between the two subpopulations and the populations are still experiencing gene flow when compared to the reference range from zero to one, where values closer to zero indicate low genetic differentiation.

The AMOVA indicates that the results are not significant, failing to reject the null hypothesis that there is no genetic differentiation among Texas and Louisiana subpopulations of F. maletae.

Source of Sum of Variance Percentage FST P-Value Variation Squares Components Variation Among 4.892 0.08191 1.01384 0.1014 0.47 Populations Among 142.639 -4.70323 -58.21310 Individuals Within Populations Within 579.500 12.70066 157.19927 Individuals Total 727.030 8.07934

Table 1. AMOVA results among populations, individuals within populations, and within individuals for Texas and Louisiana subpopulations of Faxonius maletae (average over

41 loci).

CHAPTER FOUR: DISCUSSION

The population dynamic data produced a linear relationship between length and weight for both males and females. As length increases, weight also increases. My study indicates that there may be sex-differentiated sampling method bias for F. maletae. Prior research indicated that the type of bait as well as the sampling method utilized can determine the sex and size of the crayfish captured (Price and Welch 2009). As with our study, using fried chicken in minnow traps attracted solely males and it is possible that males are more mobile than females. Females were only captured utilizing active methods of electroshocking and dip-netting. This could also indicate that females do not forage as far as males or do not take as many risks because of the rearing of young.

Trapping also attracted larger individuals and was biased toward Form I males which corresponds with other studies indicating this type of bias exists for trapping (Price and

Welch 2009). The water level was also shallow in the majority of the sampling sites and

I captured smaller crayfish in shallow water. Males captured in deeper water were larger than those captured in shallow water, which corresponds with prior research findings in manipulated experiments (Flinders and Magoulick 2007).

Several areas of Bayou Santabarb have been dammed to create cattle ponds, decreasing water flow and increasing habitat fragmentation in Louisiana (Walls 1985).

This population is also subject to heavy sedimentation increases that change stream bottom composition as a result of logging (Williams et al. 2005). The Peason Ridge

Training Area sites seem to be affected heavily by the military practices performed in the streams such as removal of debris, leaf and log litter, as well as construction rock

28 removal. As I hypothesize that F. maletae prefer rocky, gravel bottoms, this action may be influencing local populations within the military base boundary. The increased introduction of sediment and the removal of stream substrates such as rocks will only further habitat degradation, contributing to F. maletae population declines.

The utilization of Next Generation Sequencing (RAD-Seq) for ecological population genetics assists with genome-scale genetic diversity analyses of populations

(Davey and Blaxter 2011). These data elucidate whether species are phylogenetically, evolutionarily, or genetically differentiated from one another. The phylogenetic analyses indicated that the Texas and Louisiana populations are not separated in the phylogenetic tree, instead they are intermingled which is representative of one common ancestor. The inferred ancestry model of K=1 from STRUCTURE indicated that the two populations are not genetically differentiated. The confirmation that these two subpopulations are not separate species is also supported by the AMOVA results.

I have formulated hypotheses surrounding why the two subpopulations are still genetically the same although they are thought to be geographically isolated. The first hypothesis is that the Texas and Louisiana subpopulations are not actually geographically separated as they were once thought to be, and they exist outside of the two ranges they have been found within to date. Historically, a log jam termed the Great Raft was responsible for clogging the Red and Atchafalaya Rivers which led to the formation of the “Great Raft Lakes” such as Caddo Lake in Texas. This hypothesis is based on this historical formation that created a division between the two subpopulations of F. maletae that were geographically isolated as a result. This structure was removed by Henry

Miller Shreve up to 1838 and his successors continued the work later. This removal

29 removed navigation impediments of the Red River and the city of Shreveport is named after him (Holbrook 2007). This raft formed again later, but it was located further up river than the prior raft and extended up to the Arkansas state line from Louisiana. The second raft was removed in 1873 by Lieutenant Eugene Woodruff (Bagur 2001;

Holbrook 2007). The second hypothesis is that the two subpopulations are actually geographically isolated from one another and they have not been separated long enough to see effects of restricted gene flow. To test either hypothesis, more sampling throughout Louisiana and Texas tributaries falling between the two already identified subpopulations would be necessary.

Performing field sampling allowed for population dynamic and size-class distribution estimates. The resulting RAD-Seq analyses provided a SNP dataset used to perform phylogenetic analyses such as a maximum likelihood approach with RAxML, a

Bayesian clustering approach with STRUCTURE, and AMOVA analyses. Upon review of all data, I conclude that the two populations are not genetically isolated or have not been isolated long enough for allopatric speciation to occur as a result of geographic isolation (Hoskin et al. 2005).

Prior research performed by Brown (2017) concluded that the Texas and

Louisiana subpopulations of Faxonius maletae were potentially different species based on mtDNA analyses. Mitochondrial DNA results indicated that the two subpopulations were genetically differentiated and were undergoing divergence at the COI and 16S gene segments. The COI gene percent sequence divergence was 5.91% and the 16S gene percent sequence divergence was 2.37% between the subpopulations (Brown 2017).

Mitochondrial DNA exhibits a rapid mutation rate, giving the appearance that a species is

30 rapidly evolving, or undergoing speciation. It is therefore necessary to use nDNA in combination with mtDNA to more conclusively explain evolutionary relationships among and between species.

One explanation for the genetic similarity in nDNA between the Louisiana and

Texas subpopulations of Faxonius maletae, if these two subpopulations are indeed geographically isolated, is that they may be at the early stages of speciation. Another explanation is that they are not actually isolated and gene flow is still occurring. As mtDNA is inherited from the mother only, or uniparentally inherited, it is expected that genes be highly conserved, yet a higher mutation rate is present in mtDNA (Gissi et al.

2008) as opposed to nDNA which is inherited from both the mother and father

(Ladoukakis and Zouros 2017). Because of the high mutation rate in mtDNA, nDNA analysis is necessary to address speciation questions because if mtDNA is the only aspect of the genome that is assessed, all individuals would be assumed to be genetically differentiated and thus be categorized as different species.

The discrepancy between mtDNA and nDNA phylogenies has revealed that histories based on mtDNA alone can be extensively misleading when determining rate of speciation in genera and the status of a species (Shaw 2002; Wiens et al. 2009).

Mitochondrial DNA analyses showed low mtDNA divergence among species; whereas, nDNA analyses show deep divergences (Wiens et al. 2009). This discrepancy has been shown in other species such as turtles (Emydidae), Hawaiian crickets (Laupala), lobster

(Thenus), freshwater crab (Aegla neuquensis), octopus vulgaris, and butterflies

(Polygonia) (Shaw 2002; Wahlberg et al. 2009; Weins et al. 2009; Barber et al. 2012;

Jeena et al. 2015; Amor et al. 2019). The conclusion is that the two subpopulations are

31 not genetically differentiated in Texas and Louisiana based on nuclear DNA genomic analysis. I feel the weight of evidence, given new data provided by nDNA results, indicates that the Louisiana and Texas sub-populations should be considered to be one species.

Management implications of my research indicate that Faxonius maletae needs to be state listed in Texas and Louisiana, effective immediately, to begin conservation of a declining species. As the species has a limited native geographic range and is potentially experiencing habitat fragmentation, a decline in suitable habitat is likely to continue. The species should be listed federally as threatened or endangered because of these documented declines across its native range.

REFERENCES

Adams, S., G.A. Schuster, and C.A. Taylor. 2010. Orconectes maletae. The IUCN Red

List of Threatened Species 2010:

http://dx.doi.org/10.2305/IUCN.UK.20103.RLTS.T15433A4584468.en.

Amor, M.D., S.R. Doyle, M.D. Norman, A. Roura, N.E. Hall, A.J. Robinson, T.S.

Leite, and J.M. Strugnell. 2019. Genome-wide sequencing uncovers cryptic

diversity and mito-nuclear discordance in the Octopus vulgaris species

complex. Review round 2: Scientific Reports.

Bagur, J.D. 2001. A History of Navigation on Cypress Bayou and the Lakes. University

of North Texas Press. Denton, TX.

Barber, B.R., J. Xu, M. Perez-Losado, C.G. Jara, and K.A. Crandall. 2012. Conflicting

Evolutionary Patterns Due to Mitochondrial Introgression and Multilocus

Phylogeography of the Patagonian Freshwater Crab Aegla neuquensis. Plos One

7(6)

Berger, S.A., D. Krompass, and A. Stamatakis. 2011. Performance, Accuracy, and Web

Server for Evolutionary Placement of Short Sequence Reads under Maximum

Likelihood. Systematic Biology 60(3):291—302.

Blanco‐Bercial, L. and A. Bucklin. 2016. New view of population genetics of

zooplankton: RAD‐seq analysis reveals population structure of the North Atlantic

planktonic copepod Centropages typicus. Molecular Ecology 25(7):1566—1580.

Brown, L.B. 2017. Phylogenetic and ecological analysis of two populations of the

Kisatchie Painted Crayfish, Orconectes maletae (Decapoda: Cambaridae). M.S.

Thesis, The University of Texas at Tyler.

Cariou, M., L. Duret, and S. Charlat. 2013. Is RAD‐seq suitable for phylogenetic

inference? An in silico assessment and optimization. Ecology and Evolution 3(4):

846—852.

Clifford, H.F. 1991. Aquatic invertebrates of Alberta. Edmonton, Alberta, Canada:

University of Alberta Press: 180.

Crandall, K.A. and J.F. Fitzpatrick Jr. 1996. Crayfish molecular systematics: Using a

combination of procedures to estimate phylogeny. Systematic Biology 45:1–26.

Crandall, K.A., D.J. Harris, and J.W. Fetzner Jr. 2000. The monophyletic origin of

freshwater crayfishes estimated from nuclear and mitochondrial DNA sequences.

Proceedings of the Royal Society of London B 267:1679–1686.

Crandall, K.A. and J.E. Buhay. 2007. Global diversity of crayfish (Astacidae,

Cambaridae, and Parastacidae—Decapoda) in freshwater. Hydrobiologia

595:295—301.

Crandall, K.A. and S. De Grave. 2017. An updated classification of the freshwater

crayfishes (Decapoda: Astacidae) of the world, with a complete species list.

Journal of Crustacean Biology 37: 615—653.

Davey, J.W. and M.L. Baxter. 2011. RadSeq: next-generation population genetics.

Briefings in Functional Genomics 9:416—423.

Desjardins, P., J.B. Hansen, and M. Allen. 2009. Microvolume Protein Concentration

Determination using the NanoDrop 2000c Spectrophotometer. Journal of Visual

Experiments 33:1610.

Durand, J.B. 1960. Limb Regeneration and Endocrine Activity in the Crayfish. The

Biological Bulletin 118:250—261.

Excoffier, L. and H.E.L. Lischer. 2010. Arlequin Suite Version 3.5: A new series of

programs to perform population genetics analyses under Linux and Windows.

Molecular Ecology Resources 10:564—567.

Faxon, W. 1884. Descriptions of new species of Cambarus, to which is added a

synonymical list of the known species of Cambarus and Astacus. Proceedings of

the American Academy of Arts and Sciences 20:107—158.

Faxon, W. 1898. Observations on the Astacidae in The United States National Museum

and in The Museum of Comparative Zoology, with Descriptions of New

Species. Proceedings of The United States National Museum 22:643—694.

Fetzner, J.W., Jr. 1996. Biochemical systematics and evolution of the crayfish genus

Orconectes (Decapoda: Cambaridae). Journal of Crustacean Biology 16:111–141.

Flinders, C.A. and D.D. Magoulick. 2007. Effects of depth and crayfish size on predation

risk and foraging profitability of a lotic crayfish. Journal of the North American

Benthological Society 26:767—778.

Gissi, C., F. Iannelli, and G. Pesole. 2008. Evolution of the mitochondrial genome of

Metazoa as exemplified by comparison of congeneric species. Heredity

101(4):301—320.

Gontcharov, A.A., B. Marin, and M. Melkonian. 2004. Are combined analyses better

than single gene phylogenies? A case study using SSU rDNA and rbcL sequence

comparisons in the Zygnematophyceae (Streptophyta). Molecular Biology and

Evolution 21(3):612—624.

Hedges, S.B. 2002. The origin of evolution of model organisms. Nature Reviews

Genetics 3:612—624.

Hobbs, H.H. 1989. An illustrated checklist of the American crayfishes (Decapoda,

Astacidae, Cambaridae, and Parastacidae). Smithsonian Contributions to Zoology.

480.

Hobbs, H.H. and D.M. Lodge 2010. Ecology and Classification of North American

freshwater invertebrates. Elsevier Science and Technology Books.

Holbrook, S.H. 2007. Lost Men of American History. Creative Media Partners,

LLC. 404.

Hoskin, C.J., M. Higgie, K.R. McDonald, and C. Moritz. 2005. Reinforcement drives

rapid allopatric speciation. Nature 437:1353—1356.

Jeena, N.S., A. Gopalakrishnan, E.V. Radhakrishnan, J.K. Kizhakudan, V.S. Basheer,

P.K. Asokan, and J.K. Jena. 2016. Molecular phylogeny of commercially

important lobster species from Indian coast inferred from mitochondrial and

nuclear DNA sequences. Mitochondrial DNA Part A 27(4).

Johnson, D.P. 2010. Four new crayfishes (Decapoda: Cambaridae) of the genus

Orconectes from Texas. Zootaxa 2626: 1—45.

Kichler, C.E. 1984. Evaluating Survival of the Crayfish Orconectes nais Exposed to

Hypoxic Winter Conditions. M.S. Thesis. Kansas State University.

Kimmel, W.G. and D.G. Argent. 2016. Community concordance between fishes and

benthic macroinvertebrates among adventitious and ordinate tributaries or a major

river system. Ecological Indicators 70:15—22.

Klecka, J. and D. Boukal. 2014. The effect of habitat structure on prey mortality depends

on predator and prey microhabitat use. Oecologia 176(1):183—191.

Ladoukakis, E.D. and E. Zourous. 2017. Evolution and inheritance of animal

mitochondrial DNA: rules and exceptions. Journal of Biological Research 24(2).

Maddison, W.P. and L.L. Knowles. 2006. Inferring phylogeny despite incomplete lineage

sorting. Systematic Biology 55:21—30.

Mathews, L.M., L. Adams, E. Anderson, M. Basile, E. Gottardi, and M.A. Buckholt.

2008. Genetic and morphological evidence for substantial hidden biodiversity in a

freshwater crayfish species complex. Molecular Phylogenetics and Evolution

48:126—135.

Meador, M.R., J.P. McIntyre, and K.H. Pollock. 2011. Assessing the efficacy of single-

pass backpack electrofishing to characterize fish community structure.

Transactions of the American Fisheries Society 132:39—46.

Munasinghe, D.H.N., N. P. Murphy, and C. M. Austin. 2003. Utility of mitochondrial

DNA sequences from four gene regions for systematic studies of Australian

freshwater crayfish of the genus Cherax (Decapoda: Parastacidae). Journal of

Crustacean Biology 23: 402—417.

Narum, S.R., C.A. Buerkle, J.W. Davey, M.R. Miller, and P.A. Hohenlohe. 2013.

Genotyping‐by‐sequencing in ecological and conservation genomics. Molecular

Ecology 22(11):2841—2847.

Nowicki, P., T. Tirelli, R.M. Sartor, F. Bona, and D. Pessani. 2008. Monitoring crayfish

using a mark-recapture method: potentials, recommendations, and limitations.

Biodiversity and Conservation 17:3513—3530.

Ogilvie, H.A., J. Heled, D. Xie, and A.J. Drummond. 2016. Computational performance

and statistical accuracy of* BEAST and comparisons with other methods.

Systematic Biology 118.

Pamilo, P. and M. Nei. 1988. Relationships between gene trees and species trees.

Molecular Biology and Evolution 5:568—583.

Pehnke, L. and A. Pyae. 2012. Reproductive morphology and sperm depletion in

crayfish. Worchester Polytechnic Institute. B.S. Project, The University of Texas

at Tyler.

Penn, G.H., Jr. 1952. A new crawfish of the Virilis Section of the genus Orconectes

(Decapoda, Astacidae). National History Miscellaneous 109:1—7.

Philippe, H., H. Brinkmann, D.V. Lavrov, D.T.J. Littlewood, M. Manuel, G. Wörheide,

and D. Baurain. 2011. Resolving Difficult Phylogenetic Questions: Why More

Sequences Are Not Enough. Plos Biology 9(3):1—10.

Porras-Hurtado, L., Y. Ruiz, C. Santos, C. Phillips, A. Carracedo, and M.V. Lareu. 2013.

An overview of STRUCTURE: applications, parameter settings, and

supporting software. Frontiers in Genetics 4:98.

Price, J.E. and S.M. Welch. 2009. Semi-quantitative methods for crayfish sampling: sex,

size, and habitat bias. Journal of Crustacean Biology 29(2):108—216.

Pritchard, J.K., M. Stephens, N. A. Rosenberg, and P. Donnelly. 2000. Association

mapping in structured populations. American Journal of Human Genetics 67:170—

181.

RStudio Team. 2016. RStudio: Integrated Development for R. RStudio, Inc. Boston, MA.

Version 1.0.153. URL http://www.rstudio.com/.

Rabeni, C.F., K.J. Collier, S.M. Parkyn, and B.J. Hicks. 1997. Evaluating techniques for

sampling stream crayfish (Paranephrops planifrons). New Zealand Journal of

Marine and Freshwater Research 31:693—700.

Rohlf, F.J. and R.R. Sokal. 1995. Statistical Tables. Third Edition. W.H. Freeman and

Company. New York.

Shaw, K.L. 2002. Conflict between nuclear and mitochondrial DNA phylogenies of a

recent species radiation: What mtDNA reveals and conceals about modes of

speciation in Hawaiian crickets. Proceedings of the National Society of Sciences of

the United States of America 99(25):16122—16127.

Simbolo, M., M. Gottardi, V. Corbo, M. Fassan, A. Mafficini, G. Malpeli, R.T. Lawlor,

and A. Scarpa. 2013. DNA Qualification Workflow for Next Generation

Sequencing of Histopathological Samples. PLOS One 8(6):1—8.

Somers, K.M. and D.P.M Stechey. 1986. Variable Trappability of Crayfish Associated

with Bait Type, Water Temperature and Lunar Phase. American Midland Naturalist

116:34—44.

Spinks, P.Q., R.C. Thomson, G.B. Pauly, C.E. Newman, G. Mount, and H.B. Shaffer.

2013. Misleading phylogenetic inferences based on single-exemplar sampling in

the turtle genus Pseudemys. Molecular Phylogenetics and Evolution 68(2):269—

281.

Stamatakis, A. 2014. RAxML version 8: a tool for phylogenetic analysis and post-

analysis-of large phylogenies. Bioinformatics 30:1312—1313.

Taylor, C.A. and J.H. Knouft. 2006. Historical influences on genital morphology among

sympatric species: gonopod evolution and reproductive isolation in the crayfish

genus Orconectes (Cambaridae). Biological Journal of the Linnean Society 89:1—

12.

Taylor, C.A., G.A. Schuster, J.E. Cooper, R.J. DiStefano, A.G. Eversole, P. Hamr, H.H.

Hobbs, H.W. Robison, C.E. Skelton, and R.F. Thoma. 2007. A reassessment of the

conservation status of crayfishes of the United States and Canada after 10+ years of

increased awareness. Fisheries 32:372—389.

Wahlberg, N., E. Weingartner, A.D. Warren, and S. Nylin. 2009. Timing major conflict

between mitochondrial and nuclear genes in species relationships of Polygonia

butterflies (Nymphalidae: Nymphalini). BMC Evolutionary Biology 9(92).

Walls, J.G. 1972. Three new crayfishes related to Orconectes difficilis (Faxon)

(Decapoda: Astacidae). Proceedings of the Biology Society of Washington

84:449—458.

Walls, J.G. 1985. Distribution and natural history of the crawfish Orconectes difficilis

(Decapoda: Astacidae) in Louisiana. Southwestern Naturalist 30:189—194.

Walls, J.G. 2005. Crawfishes from the Kisatchie Ranger District, Kisatchie National

Forest, Natchitoches Parish, Louisiana. Presented in fulfillment of a

contract with Mr. David Byrd, Fisheries Program Manager, KNF, Pineville, LA.

Weins, J.J., C.A. Kuczynski, and P.R. Stephens. 2009. Discordant mitochondrial and

nuclear gene phylogenies in emydid turtles: implications for speciation and

conservation. Biological Journal of the Linnean Society 99:445—461.

White, D.J., J.N. Wolff, M. Pierson, N.J. Gemmell. 2008. Revealing the hidden

complexity of mtDNA inheritance. Molecular Ecology 17(23):4925—4942.

Williams, L.R., T.H. Bonner, J.D. Hudson, M.G. Williams, T.R. Leavy, and C.S.

Williams. 2005. Interactive effects of environmental variability and military

training on stream biota of three headwater drainages in western Louisiana.

Transactions of the American Fisheries Society 134:192—206.

Williams, L.R., M.G. Williams, J. Banta, J. Hernandez, L. Brown, and J. Placyk. 2014.

Ecological niche modeling and field surveys for the Kisatchie painted crayfish,

Orconectes maletae. USFWS SWG Contract 447170.

Wright, S. 1965. The interpretation of population structure by F-statistics with special

regard to systems of mating. Evolution 19:395—420.

APPENDIX A

Table 2: Texas historical sites for Faxonius maletae sampled in 2017—2018.

Site Label County Latitude Longitude Dates Sampled Sample Labels T1 Upshur 32.77541 -94.94578 6/25/2018 None T2 Upshur 32.79811 -95.04985 6/25/2018 None T3 Gregg 32.67281 -94.75155 6/25/2018 None T4 Harrison 32.62358 -94.67286 6/25/2018 None T11 Marion 32.78889 -94.51634 6/25/2018 None T13 Marion 32.7497 -94.49978 6/25/2018 None T16 Marion 32.75633 -94.34306 6/25/2018 None T19 Titus 33.02177 -94.88128 6/24/2018, T19-1—T19-3 10/4/2018, 10/5/2018 T20 Titus 33.07185 -94.96546 6/25/2018, T20-1—T20-3 10/5/2018 T23 Franklin 33.0511 -95.14247 6/25/2018 None

APPENDIX A: CONTINUED

Table 3: Louisiana historical sites for Faxonius maletae sampled in 2017—2018.

Site County Latitude Longitude Dates Sample Label Sampled Labels L5 Natchitoches 31.5636667 -93.2401667 6/21/2018 L5-1—L5-8 Parish L6 Natchitoches 31.5362833 -93.2067 6/21/2018 L6-1 Parish L7 Natchitoches 31.4847833 -93.13845 6/21/2018 L7-1—L7-3 Parish L8 Natchitoches 31.445833 -93.096806 6/21/2018 None Parish L9 Natchitoches 31.422683 -93.170867 6/21/2018 None Parish L10 Natchitoches 31.408933 -93.171083 6/21/2018 None Parish L11 Natchitoches 31.3952833 -93.0636 6/20/2018, L11-1— Parish 6/21/2018 L11-11 L12 Natchitoches 31.3986833 -93.0921167 6/24/2018 L12-1 Parish L13 Natchitoches 31.398950 -93.106750 6/21/2018 None Parish L14 Natchitoches 31.4318333 -92.94245 6/21/2018 L14-1 Parish L26 Natchitoches 31.410108 -93.169194 6/21/2018 None Parish

APPENDIX A: CONTINUED

Table 4: Sites sampled for Faxonius maletae on Fort Polk military base in Leesville, LA in 2017—2018.

Site Label County Latitude Longitude Dates Sampled Sample Labels LC Natchitoches 31.3881853 -93.2558782 10/2017, 04/2018— LC1—LC12 Parish 06/2018, 09/2018, 10/2018 TC Natchitoches 31.3883155 -93.1856801 10/2017, 04/2018— TC1—TC3 Parish 06/2018, 09/2018, 10/2018 SC Natchitoches 31.3783322 -93.2464596 10/2017, 04/2018— SC1 Parish 06/2018, 09/2018, 10/2018 OC Natchitoches 31.3854161 -93.1974174 10/2017, 04/2018— OC1, OC2 Parish 06/2018, 09/2018, 10/2018

Table 5: Average Maximum Log Likelihood from STRUCTURE results.

K Average Maximum Log Likelihood 1 -2239.466667 2 -2241.066667 3 -2240.733333 4 -2240.1 5 -2239.6 6 -2240.433333 7 -2241.3 8 -2241.4 9 -2239.933333 10 -2240.133333 11 -2239.033333 12 -2240.133333 13 -2236.933333 14 -2239.933333 15