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Extensive Phylogeographic and Morphological Diversity in Diporiphora nobbi (Agamidae)
Leads to a Taxonomic Review and a New Species Description
Author(s) :Danielle L. Edwards and Jane Melville
Source: Journal of Herpetology, 45(4):530-546. 2011.
Published By: The Society for the Study of Amphibians and Reptiles
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Extensive Phylogeographic and Morphological Diversity in Diporiphora nobbi (Agamidae)
Leads to a Taxonomic Review and a New Species Description
DANIELLE L. EDWARDS
AND JANE MELVILLE
Department of Sciences, GPO Box 666, Museum Victoria, Melbourne, VIC 3001, Australia
ABSTRACT.—Morphological and molecular information is invaluable in the description of cryptic diversity and the evolutionary processes
driving diversification within closely related species that exhibit morphological homoplasy. We present a distribution-wide data set consisting
of both molecular and morphological information, providing a taxonomic revision of the Diporiphora nobbi species group, and develop
preliminary hypotheses regarding the evolutionary history of D. nobbi. We show deep molecular divergence between D. nobbi and a newly
described sister lineage associated with divergence in meristic characters. Our molecular data also show large divergences among subclades
within nominate D. nobbi associated with different habitats rather than specific biogeographc barriers. We further discuss potential
diversification mechanisms within the D. nobbi species group.
Integration of morphological and molecular data has become
a powerful tool for resolving taxonomic issues, diversification
mechanisms, and biogeographic pattens, particularly for
groups with traditionally uncertain or difficult taxonomic
reconstruction. The Australian agamid lizards are one such
group where both higher level, inter- and intrageneric
taxonomic relationships have been difficult to resolve due to
a lack of diagnostic morphological characters and high levels of
intraspecific phenotypic variation. Molecular techniques have
proven invaluable in resolving the higher level systematics of
the group (Macey et al., 2000; Schulte et al., 2003; Hugall and
Lee, 2004; Hugall et al., 2008) that remained uncertain under a
purely morphological framework. Molecular studies also have
shown that the Australian agamids form a single evolutionary
lineage (Macey et al., 2000), known as the subfamily Amphi-
bolurinae, that diversified across Australia’s arid and semiarid
regions during the Miocene in response to an increasingly dry
environment (Hugall et al., 2008).
More recent intrageneric molecular work has demonstrated
that there is great variation in the level of morphological
diversity within evolutionary lineages of the Amphibolurinae.
One well-studied genus within Amphibolurinae, Ctenophorus,
has shown dramatic diversity in morphological evolution
between closely related species (Melville et al., 2001). However,
the opposite has been found in other genera, with widespread
‘‘species’’ being found to be cryptic species complexes
(Melville et al., 2008; Shoo et al., 2008), some of which display
morphological stasis rather than disparity (Smith et al., 2011).
Such variation in the level of morphological diversity within
lineages reinforces the importance of integrating molecular and
morphological data when undertaking taxonomic reviews in
the Australian agamid lizards.
Diporiphora is one of the more species-rich genera within
Amphibolurinae and contains many lineages for which
taxonomic revision is needed. Diporiphora nobbi, a recent
addition to this genus after molecular revision (Schulte et al.,
2003; Hugall and Lee, 2004; Hugall et al., 2008), has a
distribution that encompasses the majority of eastern Australia.
The huge distribution and the extensive morphological
variation seen in this species suggest that an investigation of
its phylogeographic variation and a taxonomic review is
Witten (1972) originally described D. nobbi as a member of
the Amphibolurus muricatus species group from material
collected across New South Wales (NSW) and southeastern
Queensland (QLD), assigning the species name Amphibolurus
nobbi. Although affinities between A. nobbi and Diporiphora
were not directly analyzed in the original description of the
species by Witten (1972), Witten did in fact note the
‘‘superficial resemblance’’ between A. nobbi and Diporiphora.
However, Witten (1972) considered Amphibolurus muricatus the
sister species to the new taxon based on the presence of femoral
pores in A. nobbi, which are generally not present in Diporiphora
species. Later, Greer (1989) noted that color patterns in A. nobbi
were similar to those of other species of Diporiphora, rather than
to Amphibolurus, with a pink or rose flush to the base of the tail
and yellowish sides. Molecular analyses have since revised the
taxonomy of the species to show that A. nobbi was in fact nested
within Diporiphora (Schulte et al., 2003; Hugall and Lee, 2004;
Hugall et al., 2008). These molecular analyses have included
multiple loci of both mitochondrial DNA (mtDNA) and
nuclear DNA and are supported by several phylogenetic
studies undertaken on Amphibolurinae in recent times (Schulte
et al., 2003; Hugall and Lee, 2004; Hugall et al., 2008). Thus, we
refer to this species as D. nobbi.
There are currently two described subspecies within D. nobbi,
D. n. nobbi and D. n. coggeri, both described from the cool
temperate forests and upland areas of the New England
Tablelands (Fig. 1). Wells and Wellington (1985) described
another species from within the range of D. nobbi and ascribed
this species to its own genus with the name Wittenagama
parnabyi based on a single specimen from central Queensland
in the vicinity of Alpha. However, this species was later
synonymized with D. nobbi by Shea and Sadlier (1999).
The distribution of D. nobbi covers steep environmental and
ecological gradients, from rain forest, dry woodlands, and
coastal swamplands and vegetation to cool temperate forests
and upland areas, as well as dry mallee with spinifex
woodland. This distribution also crosses many important
biogeographic barriers known to delineate population struc-
ture in its sister lineage D. australis (Edwards and Melville,
2010) and many other woodand, highland, and rain forest
species throughout QLD and NSW (Schneider et al., 1998;
Moritz et al., 2000; Dolman and Moritz, 2006; Moussalli et al.,
2009). Diporiphora nobbi shows a large amount of geographic
variation in body size, morphology, and male secondary sexual
coloration at the tail base (e.g., often displaying a purple, pink,
or red flush; Cogger, 2000), with some forms showing stark
body coloration (e.g., the Alpha form having a canary yellow
body in displaying males; pers. obs.).
To clarify the taxonomic status of D. nobbi, we compiled a
multilocus molecular data set in combination with morpho-
logical data collected from museum voucher specimens from
across the distribution. Specifically, we sought to test whether
D. n. nobbi,D. n. coggeri (Witten, 1972), and W. parnabyi (Wells
Corresponding Author. Present address: 119A Environmental
Science Center, Department of Ecology and Evolutionary Biology, Yale
University, New Haven, CT 06520-8106 USA. E-mail: Dan.Edwards@
Journal of Herpetology, Vol. 45, No. 4, pp. 530–546, 2011
Copyright 2011 Society for the Study of Amphibians and Reptiles
and Wellington, 1985) could be ascribed species status.
Providing a complete assessment of molecular and morpho-
logical variation in D. nobbi, we contribute an important
advance in our understanding of the biogeographic history
and taxonomy of eastern Australian vertebrate fauna.
MATERIALS AND METHODS
Taxonomic Sampling.—Sampling effort focused on gathering
tissue samples from across the distributions of both nominate
subspecies in the D. nobbi species group—D. n. nobbi and D. n.
coggeri. This was accomplished using a combination of tissues
deposited in Australian museums and fresh tissue collections.
In total, 90 individuals were sampled from across the species
distribution, including the distributions of both nominal
subspecies, with one to five animals per site (Fig. 1a; for
details on each sample, see Appendix 1). For fresh field
collections two specimens per site were deposited in the
Museum Victoria Collection (liver tissue); the remaining
animals from each site were nonlethally sampled (tail-tips)
and released at the point of capture. Specimens and tissues
collected fresh have been listed in Appendix 1 (MVD and MVZ
Museum samples were obtained from the Australian
Museum Tissue Collection, Australian Biological Tissue Col-
lection (South Australian Museum), and the Queensland
Museum Tissue Collection. Morphological measurements were
taken from all suitable specimens for which molecular data
were available. In addition, to encompass the morphological
variation across the group and determine appropriate species
assignment, specimens representing gaps in tissue collections
and potential species boundaries were morphologically mea-
sured in conjunction with holotypes or paratypes (For
distribution of material examined, see Fig. 1b; for details of
material examined, see Appendix 2). Paratypes were included
for both nominal subspecies within the D. nobbi species
complex (D. n. nobbi and D. n. coggeri) in addition to the
holotype for W. parnabyi (Wells and Wellington, 1985), which
was synonymized with D. nobbi by Shea and Sadlier (1999).
Outgroup samples for mtDNA analyses were selected
from across Diporiphora and associated genera (D. australis,
GU556007; D. bilineata, AF128473; A. nobbi, AY132999; D.
albilabris, AY133003; D. arnhemenica, AY133004; Caimanops
amphiboluroides, AF128472; D. magna, AY133009; D. winneckii,
AY133012; and A. muricatus, AF128468). For specific informa-
tion on each of these ND2 sequences, refer to Melville et al.
(2001), Schulte et al. (2003), and Edwards and Melville (2010).
For recombination activating gene-1 (RAG1) analyses, out-
groups were obtained from previously published sequences
from across Diporiphora and associated genera (Appendix 3). In
addition, sequence data were obtained for D. australis
(MVD74106; vicinity of Charters Towers [20u12957.20E 146u
14943.40S]; GenBank #JN815263) for RAG1 to supplement
these sequence data.
FIG. 1. Map of sampling locations for tissues used for genetic analysis (a, N) and material examined for morphological analyses (b, &). The type
localities for each of the nominate subspecies within Diporiphora nobbi are shown (D. n. nobbi, open black and gray star; D. n. coggeri, solid black star),
as is the type locality for the previously synonymized Wittenagama parnabyi (open gray and white star). Map of Australia is inset.
SYSTEMATICS OF THE DIPORIPHORA NOBBI SPECIES COMPLEX 531
Molecular Data.—Genomic DNA was extracted from the tail
and liver samples by using a modified chloroform method,
suspended in Tris-EDTA buffer, and stored at 220uC (for
details, see Shoo et al., 2008). For all individuals, we targeted an
,1,400-bp fragment of mtDNA incorporating the entire
protein-coding gene NADH dehydrogenase subunit 2 (ND2)
and the genes encoding tRNA
to the beginning of the protein-coding gene
subunit I of cytochrome coxidase. For a subset of these
animals, we sequenced an ,1,400-bp fragment of nuclear DNA
incorporating a portion of the RAG1 exon to ensure resulting
relationships could be confirmed across multiple mtDNA and
Targeted DNA was amplified using a touch-down polymer-
ase chain reaction (PCR) profile (94uC for 5 min, 13;94uC for
30 sec, 70–45uC (decreasing in 5uC increments) for 20 sec, 72uC
for 90 sec, 23;94uC for 30 sec, 40uC for 30 sec, 72uC for 45 sec,
403;72uC for 4 min, 13;4uC, hold. Primers used to amplify
ND2 were Metf-1 (59-AAGCAGTTGGGCCCATRCC-39, com-
plement of H4419b) (Macey et al., 2000) and H5934 (59-
AGRGTGCCAATGTCTTTGTGRTT-39) (Macey et al., 1997) or
CO1r.aga (59-ACRGTTCCRATRTCTTTRTGRTT-39) (Macey et
al., 2000). Primers used to amplify RAG1 were JRAG1f.1 (59-
CAAAGTGAGACSACTTGGAAAGCC-39) and JRAG1r.13 (59-
CATTTTTCAAGGGTGGTTTCCACTC-39) (Shoo et al., 2008).
Targeted fragments were amplified in 40-ml reactions consist-
ing of ,100 ng of template DNA, 4 mlof103reaction buffer,
3 mM MgCl
, 0.5 mM dNTPs, 10 pmol of each primer, and 2
units HotStart Taq polymerase (MBI Fermentas). PCR products
were purified using a SureClean PCR cleanup kit (Bioline) or
gel purified using GFX columns (GE Healthcare) and then sent
to Macrogen Inc. (Seoul, Korea) for sequencing. Internal
primers ND2f.17 (59-TGACAAAAAATTGCNCC-39) (Macey
et al., 2000) and ND2f.dip (59-AAATRATAGCCTACTCAT-39)
(Shoo et al., 2008) were used in addition to PCR primers to
obtain reliable sequence across the ND2 gene. DNA sequence
data were then edited using Sequencher 4.1.4 (Gene Codes
Corporation). Sequences were aligned individually using
ClustalX (Thompson et al., 1997). Alignments were then
checked by eye. Protein coding regions were translated using
the mammalian genetic code option in Sequencher 4.1.4 (Gene
Codes Corporation), and a clear reading frame was observed in
all ND2 sequences. Thus, sequences were assumed to be
genuine mitochondrial copies and not nuclear paralogs.
Sequences have been lodged in GenBank (Appendix 1).
Phylogenetic Analyses.—Bayesian and maximum likelihood
(ML) analyses of haplotype sequences were used to assess
overall phylogenetic structure and support for major clades in
Mr Bayes version 3.1.2 (Ronquist and Huelsenbeck, 2003) and
RAxML (Stamatakis et al., 2008) individually for both ND2 and
RAG1 data sets. Modeltest 3.7 (Posada and Crandall, 1998),
using Akaike information criterion showed that for the mtDNA
data the GTR +I+Cmodel of nucleotide substitution best fit
the data, and for the RAG1 data the TRN +I+Cmodel of
nucleotide substitution best fit the data. Bayesian and ML
analyses were conducted using these best fit models of
nucleotide substitution and partitioned according to codon
position for mtDNA data. Bayesian analyses were undertaken
using default priors for Markov chain Monte Carlo analyses in
MrBayes version 3.1.2 (Ronquist and Huelsenbeck, 2003).
Bayesian analyses consisted of four independent runs of four
chains each, sampling every 100 generations run for 10 310
generations for the ND2 data set and for 4 310
the RAG1 data set to ensure convergence. Burn-in was set at
1,000,000 and 400,000 generations for the ND2 and RAG1
analyses, respectively. Convergence of posterior probabilities
and stationarity of likelihood scores between the runs were
assessed in Tracer version 1.4 (Rambaut and Drummond,
2005). ML analyses were run for 100 bootstrap replicates.
Estimates of Divergence Times.—We estimated divergence
times for both the ND2 and RAG1 data sets separately using
the relaxed molecular clock method in the program BEAST
version 1.6.1 (Drummond and Rambaut, 2007). Initial analyses
used several fossil calibrations across Reptilia, and specifically
within the Iguania, we used an expanded dating data set as per
Shoo et al. (2008) and Melville et al. (2009), sampling across the
Reptilia by using GenBank sequences (for outgroups used and
their associated accessions, see BEAST output in Appendix 4).
Given the issues regarding overestimated divergences caused
by saturation-driven branch length truncation (Hugall et al.,
2007; Sanders et al., 2008), RAG1 and mtDNA analyses were
run separately. Analysis of the RAG1 data set used fossil
calibrations across the Iguania to estimate the divergence
between D. nobbi and D. sp. nov. Fossil calibrations included
four fossils used in previously published studies: a middle
Jurasic acrodont iguanian fossil (154–180 mya, Evans et al.,
2002), an early Miocene sceloporine (,22.8 mya , Robinson and
Van Devender, 1973) and an Chamaeleo/Rhampholeon fossil
(18 mya, Rieppel et al., 1992), and a Pliocene Phrynocephalus
fossil (5 mya, Zerova and Chkhikvadze, 1984). Specific BEAST
settings for these calibrations are as per table 1 in Melville et al.
(2009). In addition, we added a minimum age estimate for the
Physignathus lesueurii lineage (which includes that majority of
the Australian amphibolurine species, except Moloch,Chelosa-
nia, and Hypsilurus) of 20 mya (Covacevich et al., 1990) with
BEAST settings as follows: mean 51.8, SD 51.0, and zero
offset 520. The Iguania and Episquamata were held as
monophyletic and all other relationships were free to vary.
Given the difficulties in resolving the intraspecific clade
relationships within nominate D. nobbi by using the RAG1 data
and considering the difficulties with using deep calibrations
with mtDNA as mentioned above, we used a reduced
sampling approach to estimate divergence times of phylogeo-
graphic clades. For the mtDNA analysis, we used only the
Phrynocephalus and P. lesueuri fossil calibrations using the
BEAST settings described above and in Melville et al. (2009).
All relationships were allowed to vary.
Both mtDNA and nuclear analyses employed a GTR +I+C
model of evolution, by using an uncorrelated lognormal
relaxed molecular clock. A Yule Speciation Process tree prior
also was used for both analyses. RAG1 analyses were run for 15
million generations and mtDNA analyses were run for 10
million generations with burnins of 150,000 and 100,000,
respectively. All fossils are a minimum estimate of age;
therefore, we used lognormal distribution for fossil calibra-
tions. Output files were reviewed in Tracer version 1.4
(Rambaut and Drummond, 2005) to check that stationarity
had been reached, to examine the coefficient of variation, to
determine the appropriateness of a lognormal clock, and to
assess the autocorrelation of rates from ancestral to descendant
lineages (Drummond et al., 2006).
Morphological Data and Analyses.—Both morphometric and
meristic character counts were taken from 113 specimens from
across the range of D. nobbi (Fig. 1b), incorporating both
specimens with and without molecular data, holotypes of
described subspecies, and previously synonymized species
holotypes. Thirteen morphometric measurements were ob-
tained: snout–vent length (SVL), axillo–groin length (AG), head
length (HL), snout length (SL), head depth (HD), head width
(HW), nostril width (NW), interorbital width (IOW), arm
length (ArL), metacarpal length (McL), leg length (LgL),
metatarsal length (MtL), and tail length (TaL). Twenty-six
meristic characters, both continuous and categorical, also were
measured for all specimens. Continuous characters included
the following: right and left femoral pore number (RFP & LFP),
right and left preanal pore number (RPP & LPP), number of
lamellae on Finger IV (McL), number of lamellae on Toe IV
(MtL), number of infralabial scales (ILS), number of supralabial
532 D. L. EDWARDS AND J. MELVILLE
scales (SLS), tympanum width (TyW), postauricular spine
number (PAS), upper auricular spine number (UAS), second-
ary upper auricular spine number (SAUS), nuchal spine
number (NS), scapular spine number (SS). Categorical charac-
ters included the following: strength of gular fold (SGF),
strength of scapular fold (SSF), strength of paravertebral scale
keeling (PVS), regularity of paravertebral scale keeling (RPVS),
regularity of scales between PV rows (RBwPVS), nuchal spine
position (NSP), dorsolateral spines (DLS), throat color (TC),
dorso-lateral body color (DLC), and lip-scale color (LC). Refer
to Table 1 for details on categories and units and on methods of
measurement for each character.
Only adult specimens (.50 mm SVL) were measured to
avoid ontogenetic changes in morphological measurements,
and male (N564) and female (N549) animals were analyzed
separately to avoid problems associated with sexual dimor-
phism. Averages and SEM for each measurement are presented
in Table 2. We sought to differentiate species based on the
putative species to which they belonged (i.e., D. nobbi,N555
males and N542 females versus D. sp. nov., N59 males and
N57 females). There was not enough data to distinguish
whether genetic clades within D. nobbi differed substantially,
because specimens available for both morphological and
genetic analysis were limited. Morphometric and meristic data
sets were analyzed separately. All morphometric measure-
ments were log-transformed to ensure normality and then
regressed against SVL to remove any allometric effects of body
size. The residuals were then plotted against SVL to ensure the
effects of body size had been removed; these residuals were
then used for subsequent analyses. Continuous morphometric
variables were regressed against SVL to adjust for differences
in body size, and the residuals of these values were used in
Principal components analyses (PCAs) were used to deter-
mine the relevance of morphometric measurements and
meristic characters, respectively, in MYSTAT version 12 (Systat
Software Inc., Chicago, IL, USA). Initial PCAs were undertaken
to identify those variables that best diagnosed the two species
of Diporiphora (i.e., as indicated by high eigenvalues in the
PCA), these variables were used as an optimal set for further
analyses. The number of principal components (PCs) extracted
from the analysis was determined from a scree-plot analysis of
eigenvalues. Individuals were then grouped into species, and
their individual factor scores were plotted in Euclidean space.
Paired t-tests were used to test for divergence in morphometric
factor scores and meristic dimension scores between species. A
Bonferroni post-hoc test identified whether the two species
significantly differed in PC scores.
Phylogenetic Analyses.—Mitochondrial sequencing from 90
individuals yielded 60 ND2 haplotypes for a 1,378-bp fragment
including the entire protein coding gene ND2, with 376
variable sites of which 317 were parsimony informative. Total
nucleotide diversity (p) for the whole mtDNA data set was
0.05710, with p50.02615 and 0.04993 for the two major clades
associated with D. nobbi and D. sp. nov., respectively. A subset
of 24 individuals was selected from the sequenced mtDNA
individuals for nuclear phylogenetic analyses. Across these 24
individual sequences for the 1,367-bp fragment of RAG1 157 bp
was variable, 64 of which were parsimony informative, with p
50.0032 across the entire data set. Twenty of these sequences
contained at least one heterozygous site; these ambiguous sites
were coded using standard International Union of Pure and
Applied Chemistry codes.
Phylogenetic analyses of the mtDNA data uncovered two
major clades within D. nobbi. The first of these major clades was
associated with D. nobbi as a whole and contained a large
amount of genetic diversity, including six subclades broadly
associated with distinct geographic regions (Fig. 2). Animals
associated with the central NSW woodlands, western NSW
slopes and highland forests, Murray River basin mallee–
spinifex woodlands, north coastal QLD forests, and central
western QLD woodlands formed distinct clades within a
monophyletic group encompassing the majority of the known
range of D. nobbi. Relationships among the subclades within
this group were not able to be resolved using the mtDNA data
set. Reciprocally monophyletic and sister to this group was a
group of animals occurring within southeastern QLD and
coastal northern NSW.
The second major clade was associated with a previously
unidentified cryptic species from the Carnarvon Gorge area in
central western QLD. The specific relationships between this
new species, D. nobbi, and D. australis–D. bilineata were
uncertain given the mtDNA data set, and although separate
from other Diporiphora species included in the phylogeny, were
not well supported as a distinct species group (Fig. 2).
However, D. nobbi and the previously undescribed species
were clearly nested within Diporiphora as per the results of all
previous molecular studies.
Phylogenetic analyses undertaken with the RAG1 (Fig. 3)
data set could not resolve relationships among the distinct
subclades within D. nobbi. However, there was strong support
for the reciprocally monophyletic sister relationship between
D. nobbi and D. sp. nov. There was also strong support for the
D. australis species group as a sister to the D. nobbi species
complex. These data also support D. nobbi as a member of
Divergence Analyses.—The relaxed lognormal clock analysis
of the mtDNA data set (Appendix 4) produced the same in-
group topology as the phylogenetic analyses (Fig. 2). For both
the RAG1 and mtDNA analyses, there was a slight tendency
toward a positive correlation in the rate of parent to child
branches, with a covariance of 20.0121 and 0.0588, respective-
ly. However, because the 95%highest posterior distribution
(HPD; 20.17 to 0.15 and 20.1 to 0.23, respectively) included
zero, this autocorrelation was not considered significant
(Drummond et al., 2006). The coefficient of rate variation was
estimated to be 0.45 (95%HPD, 0.36–0.55) and 0.51 (95%HPD,
0.41–0.62 95%) for the RAG1 and mtDNA data sets, respec-
tively, indicating that neither data set is strictly clock-like and
that a lognormal relaxed clock is appropriate. The complete
BEAST output trees are supplied in Appendices 3 and 4,
showing the nuclear and mtDNA data sets, respectively.
Our divergence analyses (Figs. 2, 3) produced an age for
Amphibolurinae (including Physignathus cocincinus and all
other Australasian agamids) at 40.37 mya (95%HPD, 28.3–
54.2 mya) for the RAG1 data and 30.48 mya (95%HPD, 22.9–
39.9 mya) for the mtDNA data. Our RAG1 divergence estimates
are slightly elevated with broader confidence limits compared
with those reported previously for the age of Amphibolurinae
by using much larger data sets (Hugall and Lee, 2004; Hugall et
al. 2008); however, our confidence limits still overlap within the
ranges reported in these more comprehensive studies, and
some discrepancy is to be expected considering the use of a
point estimate calibration as opposed to the distribution of ages
used in our study. Our divergence estimates for the purely
Australasian Amphibolurinae (including Moloch,Hypsilurus,
and all remaining Australian agamid genera) are 29.2 mya
(95%HPD, 21.6–37.7 mya) for the RAG1 data and 24.1 mya
(95%HPD, 20.2–30.2 mya) for the mtDNA data, again
overlapping with previously published data (Hugall and Lee,
2004; Hugall et al. 2008).
Our estimates of divergence (Figs. 2, 3) suggest the age of the
Diporiphora australis–D. nobbi lineage is ,10 mya (95%HPD for
RAG1, 13.2 mya [7.7–19.2 mya]; 95%HPD for mtDNA, 10.7 mya
SYSTEMATICS OF THE DIPORIPHORA NOBBI SPECIES COMPLEX 533
TABLE 1. Morphological and morphometric characters measured for specimens of Diporiphora nobbi. Included is a detailed description of each
measurement, the abbreviation used and the type, units, or categories used to classify each measurement. DL, dorso-lateral; PAF, postauricular fold;
Type of character Character name Abbreviation Method of measurement Unit/category/type
Snout–vent length SVL Distance between the tip of the nose
and the cloaca
Axillo–groin length AG Distance between the posterior of the
forelimbs and the anterior of the
Snout length SL Distance between the anterior orbital
cavity and the tip of the nose
Head length HL Distance between the tip of the nose
and the base of the skull
Nose width NW Width of the skull between the
Interorbital width IOW Width of the skull between the
Head width HW Width of the skull at its widest
point between the occular cavity
and the base of the skull
Arm length ArL Length of the arm from the tip of
the claw on the 4th digit to where
the arm joins the body
Leg length LgL Length of the leg from the tip of the
claw on the 4th digit to where the
leg joins the body
Metacarpal length McL Distance from tip of the fourth digit
to the attachment to the hand
Metatarsal length MtL Distance from the tip of the fourth
digit to the attachment to the foot
Right femoral pores RFP No. of femoral pores on the right
Left femoral pores LFP No. of femoral pores on the left leg Count
Right preanal pores RPAP No. of preanal pores on the right
side anterior to the cloaca
Left preanal pores LPAP No. of preanal pores on the left side
anterior to the cloaca
Finger IV lamellae McL No. of lamellae on the forth digit on
Toe IV lamellae MtL No. of lamellae on the forth digit on
Infralabial scales ILS No. of scales on the upper lip Count
Supralabial scales SLS No. of scales on the lower lip Count
Tympanum width TyW Horizontal diameter of the
Postauricular spines PAS No. of spines immediately anterior to
Upper auricular spines UAS No. of spines immediately above the
Secondary upper auricular
SUAS No. of spines immediately above the
Scapular spines SS No. of scapular spines Count
Nuchal spines NS No. of nuchal spines Count
Gular fold SGF Strength of gular fold Absent/weak/strong/very strong
Scapular fold SSF Strength and length of scapular fold Absent/ weak before PV rows/ weak to
PV rows/ strong before PV rows/
strong to PV rows/ strong curves over
PVS Strength of paravertebral scale keeling Weak/moderate/strong
RPVS Regularity of PV scale keeling Even/uneven/uneven with two rows
Scales between PV rows SBwPVS Regularity of scales between PV rows
(outer and inner rows)
Both regular/outer row irregular, inner
row moderately regular/both
moderately regular/both highly
Nuchal spines (position) NSP Position and arrangement of nuchal
Absent/diagonal up from PAF no
clusters/diagonal up from PAF with
clusters/extend from DL PV scale
Dorso-lateral spines DLS Presence/absence of DL spines and
Absent/absent but some enlarged
scales/present without clusters/
present with clusters
534 D. L. EDWARDS AND J. MELVILLE
[6.9–13.9 mya]). Due to our inability to resolve the relationships
among D. nobbi,D. australis, and D. sp. nov. by using the
mtDNA data set, we estimate divergence between D. nobbi and
D. sp. nov. by using the RAG1 data set only. The RAG1
topology suggesting these two species as sister taxa is
consistent with the large degree of morphological similarity
between the two taxa and is probably a function of saturation
of mtDNA signal as opposed to suggesting uncertain relation-
ships among these members of this species group. Using the
RAG1 data, we estimate the divergence between D. nobbi and
D. sp. nov. occurred ,8.61 mya (95%HPD, 3.8–13.1 mya).
Divergences within D. nobbi firmly place intraspecific diversi-
fication within this species in the late Miocene period (3–
8 mya), with divergence between the populations of D. sp. nov.
occurring within the Pio-Pleistocene (2.25 mya; 95%HPD, 0.8–
Morphological Analyses.—PCA was used initially to reduce
the number of morphological and meristic measurements to a
fewer number of independent axes. Neither males nor females
could be distinguished based on morphometric measurements,
with these variables not strongly positively or negatively
loaded in PCA results (Table 3). In males, the first two PCs
accounted for 64.87%of the observed variation (Table 3). PC2
is negatively loaded for nuchal spine, metacarpal scale, and
metatarsal scale counts and moderately positively loaded for
right femoral pore and scapular spine counts, attributing to
17.65%of male variation (Table 3). Males with high PC1 scores
have higher numbers of femoral and preanal pores, a higher
number of spines on the head and scapular regions, irregular
scalation between the paravertebral scale rows, a low number
of metatarsal scales, and dark throat coloration, accounting for
47.22%of the variation in male characters. Males with high PC2
scores have fewer nuchal spines, metacarpal scales, and
metatarsal scales, with a moderate number of right femoral
pores and scapular spines, accounting for 17.65%of male
variation. Using PC1 scores, D. sp. nov. (PC1 51.53 60.446;
N59) exhibits significantly higher PC1 scores (t527.12,
df 551, P50.000) than D. nobbi (PC1 520.310 60.755; N5
44). For PC2 scores, D. sp. nov. individuals are slightly but
significantly higher (t522.67, df 551, P50.02; PC2 50.685
60.849) than D. nobbi (PC2 520.017 60.937). Morphological
divergences between the two species are further illustrated
when PC1 scores are plotted against PC2 scores and the 95%
confidence kernel ellipses do not overlap (Fig. 4a).
For females the first two PCs accounted for the majority of
morphological variation (59.22%, Table 3). Females with high
Type of character Character name Abbreviation Method of measurement Unit/category/type
Throat color TC Color of scales on the throat Absent/mottling grey/mottling black/
black with white chevron/black in
gular area only/all black
Dorso-lateral color DLC Color of the body on the DL surface Absent/restricted to scapular area/
strongest in scapular area but extends
onto entire DL surface/even and
covering DL surface
Lip scale color LC Color of the lip scales Indistinct from head color/demarcated
but not distinct from head color/
demarcated and white
TABLE 1. Continued.
TABLE 2. Average (with SDs) morphometric and meristic character results between Diporiphora nobbi and D. sp. nov. Refer to Table 1 for a
description of characters measured and categories.
D. nobbi D. sp. nov. D. nobbi D. sp. nov.
SVL 65.93 (69.16) 62.43 (66.93) 68.92 (68.22) 62.34 (65.81)
AG 29.01 (64.15) 26.43 (64.01) 32.23 (65.23) 28.56 (63.93)
SL 8.26 (61.28) 8.19 (61.15) 8.21 (61.02) 7.95 (60.74)
HL 21.25 (63.40) 20.66 (62.93) 21.04 (62.25) 19.77 (61.49)
HD 10.51 (61.28) 9.84 (61.00) 10.33 (60.94) 9.68 (60.84)
NW 5.88 (60.82) 5.83 (60.81) 5.95 (60.73) 5.64 (60.47)
IOW 9.31 (61.20) 9.36 (61.14) 9.25 (60.93) 8.77 (60.97)
HW 13.92 (61.76) 13.93 (61.51) 13.97 (61.29) 13.40 (61.09)
ArL 30.13 (63.87) 29.75 (62.94) 31.13 (63.06) 29.74 (61.86)
LgL 56.08 (66.87) 54.69 (65.60) 55.93 (65.55) 54.10 (64.43)
McL 6.66 (60.90) 6.42 (60.50) 6.66 (60.85) 6.13 (60.70)
MtL 13.70 (61.49) 12.81 (61.20) 13.34 (61.48) 12.26 (61.00)
TaL 166.88 (631.16) 153.13 (623.19) 162.03 (622.45) 137.57 (615.83)
RFP 1.20 (0–4) 4.56 (4–6) 1.44 (0–3) 3.67 (2–5)
LFP 1.30 (0–3) 4.44 (4–6) 1.46 (0–3) 3.50 (2–4)
RPAP 2.55 (1–3) 4.33 (3–6) 2.73 (0–4) 3.60 (3–4)
LPAP 2.58 (1–4) 4.44 (4–6) 2.63 (0–4) 3.50 (3–4)
McS 17.38 (14–22) 16.56 (16–18) 16.64 (14–20) 16.43 (15–18)
MtS 26.85 (22–32) 24.89 (22–27) 24.93 (21–29) 24.57 (23–26)
ILS 10.33 (9–13) 11.11 (10–13) 10.57 (8–13) 10.43 (8–12)
SLS 10.26 (8–13) 11.33 (11–14) 10.29 (8–12) 10.14 (8–11)
TyW 3.05 (60.43) 3.34 (60.38) 3.03 (60.40) 3.30 (60.33)
PAS 4.16 (1–8) 5.11 (2–12) 3.74 (1–9) 4.00 (2–6)
UAS 4.95 (1–8) 6.67 (5–8) 5.17 (1–10) 6.43 (6–8)
SUAS 2.47 (0–6) 4.78 (4–6) 2.38 (0–5) 3.71 (2–5)
SS 1.27 (0–6) 3.89 (1–8) 1.67 (0–4) 2.86 (2–4)
NS 3.85 (0–10) 9.11 (5–14) 3.62 (0–9) 8.00 (6–10)
SYSTEMATICS OF THE DIPORIPHORA NOBBI SPECIES COMPLEX 535
PC1 scores have high numbers of nuchal, upper auricular and
scapular spines, higher femoral pore counts, irregular scalation
between enlarged paravertebral rows, and a random or
clustered arrangement of nuchal spines separate from PoAF
and DLS, accounting for 40.59%of female morphological
variation. PC2 accounts for 18.63%of all female morphological
variation, indicating individuals with higher numbers of
supralabial, metacarpal, and metatarsal scales, higher numbers
of upper auricular spines, and fewer scapular spines. Females
differ significantly (t525.19, df 546, P50.000) in PC1
between D. sp. nov. (1.48 60.484; N57) and D. nobbi (20.223
60.84; N541) but not PC2 (t520.19, df 546, P51.000; D.
sp. nov. PC2 50.038 60.793; D. nobbi PC2 520.039 61.039).
Incomplete morphological divergence is reflected in the
confidence ellipse overlap that is seen between D. sp. nov.
females and D. nobbi females when PC scores are plotted
against one another in Euclidean space (Fig. 4b).
Diporiphora phaeospinosa sp. nov.
Holotype.—MVD74128 collected from Bauhinia Station, QLD
Paratypes.—Males: QMJ32596, Blackdown Tablelands
(23.80uS, 149.13uE); QMJ33335, QMJ33336, QMJ34296, Black-
down Tablelands (23.80uS, 149.07uE); AMR151843, AMR151844,
Blackdown Tablelands (23.76uS, 149.10uE); MVD74129 Bauhinia
Station (25.19uS, 149.16uE); QMJ36891, Glenhaughton Station
(25.23uS, 148.95uE). Females: QMJ34294, QMJ34295, QMJ36890,
Blackdown Tablelands (23.80uS, 149.13uE); QMJ50807, Black-
down Tablelands (23.80uS, 149.10uE); AMR151842, Blackdown
Tablelands (23.79uS, 149.09uE); AMR151845, Blackdown Table-
lands (23.76uS, 149.10uE); QMJ36892, Reklau Park (23.33uS,
147.50uE); QMJ38591, Glenhaughton Station (25.14uS, 148.57uE).
Juveniles: QMJ28495, Blackdown Tableland (23.80uS, 149.13uE);
QMJ30267, QMJ38560, QMJ38590, Robinson Gorge (25.28uS,
149.15uE); QMJ38589, Glenhaughton Station (25.23uS, 148.95uE).
Diagnosis and Distinction from Other Species.—Diporiphora
phaeospinosa is similar in body size and proportion to D. nobbi;
in fact, the two species cannot be distinguished using any
single morphometric trait measured. The main feature distin-
guishing the two species is the higher number of femoral and
preanal pores and nuchal spines in D. phaeospinosa in
comparison with D. nobbi. The PCA (Fig. 4) also identified
high numbers of scapular spines, auricular spines, and dark
throat coloration in males as significant factors in distinguish-
ing the species. Another distinguishing feature is the presence
of enlarged dorso-lateral scales, a feature that is also present in
juveniles and adults and that may assist in identifying
immature individuals. Both D. nobbi and D. phaeospinosa lack
femoral pores, distinguishing them from their sympatrically
distributed sister lineage, the D. australis/bilineata species
group, which has femoral pores (Witten, 1972).
Description of Holotype.—Adult male SVL, 72.87 mm; AG,
31.85 mm; HL, 25.32 mm; SL, 9.36 mm; HD, 11.32 mm; HW,
FIG. 2. Bayesian consensus phylogram of mtDNA clades within Diporiphora nobbi and a newly identified species (D. sp. nov.), including
estimated dates for specific nodes and a distribution map of each major clade. Nodal support is shown in the form of Bayesian posterior
probabilities from 4 million generations, and estimated nodal dates are median values with 95%confidence intervals displayed in brackets from 10
million generations. Six major mtDNA clades are identified within D. nobbi. Clades are generally associated with specific regions, habitat types, or
both: SE QLD & NE NSW, solid circle; CW QLD, open triangle; N Coastal QLD, solid triangle; Murray mallee–spinifex Woodlands, solid square; W
NSW Slopes and Highlands, solid diamond; and CW NSW Woodlands, plus sign. The distribution of D. sp. nov. is shown by the cross sign. The
type localities for each of the nominate subspecies within D. nobbi are shown (D. n. nobbi, open black and gray star; D. n. coggeri, solid black star), as
is the type locality for the previously synonymized Wittenagama parnabyi (open gray and white star).
536 D. L. EDWARDS AND J. MELVILLE
15.94 mm; NW, 6.58 mm; IOW, 10.59 mm; ArL, 31.22 mm; McL,
6.59 mm; LgL, 59.79 mm; MtL, 14.08 mm; TaL, 188 mm; RFP, 4;
LFP, 4; RPP, 4; LPP, 4; McL, 18; MtL, 26; ILS, 13; SLS, 14, TyW,
3.89 mm; PAS, 8; UAS, 7; SAUS, 5; NS, 9; SS, 4.
Diporiphora phaeospinosa is a robust and large lizard com-
pared with other members of the genus Diporiphora. The head
is large and angular with a sharp snout that is wider than it is
deep. Tympanum has a series of enlarged spine rows
extending anteriorly (Fig. 5b). Nuchal spines are separate from
the paravertebral scale rows on the dorsum and extend across
the base of the head in an uneven manner. Gular folding is
absent, and the throat is heavily colored with black extending
from the chest across the entire underside of the throat.
Infralabial and supralabial scales are clearly demarcated from
the darker head coloration and are white.
Scapular folding is strong and arches up across the forearm
to contact the paravertebral scale rows. Scapular spines form a
row along the scapular fold, but they do not extend along its
entire length. Specimen has dark coloration across the entire
dorso-lateral surface similar to that observed on the throat.
Scattered enlarged scales forming spines are randomly
scattered across the entire dorsolateral surface of the body.
Specimen has faint darkish colored paravertebral stripes that
are not continuous down the extent of the body. Paravertebral
scale rows are steeply keeled and distinct from surrounding
scales and form uneven rows that are separated by unevenly
scattered keeled scales (Fig. 5f). Scales between the paraverte-
bral rows are somewhat reduced toward the distal end of the
body and base of the tail. Tail is not compressed, but rounded,
and displays a deep red flush at the base extending the length
of the tail in life.
Intraspecific Variation.—Table 2 shows that morphometric
characters within D. phaeospinosa are directly overlapping with
the intraspecific variation observed within D. nobbi. Auricular,
scapular, and nuchal spine count variation is broader in D.
phaeospinosa than D. nobbi, and although the two species
directly overlap D. phaeospinosa generally has more spines
than D. nobbi. Dorsolateral spines and keeling of paravertebral
scales are consistently present in D. phaeospinosa, with few
individuals displaying similar but not identical characters in D.
nobbi. Analysis of variation in male breeding coloration was not
possible from preserved specimens and may provide further
distinguishing characteristics between the two species. The
breadth of variation possible within D. phaespinosa may not be
accurately assessed in the current study due to a limited
number of specimens available for analysis. In addition,
variation as it is currently defined within D. nobbi may be
altered with further revision of taxonomic relationships among
the divergent clades within D. nobbi.
Distribution.—The distribution of D. phaeospinosa is currently
limited to four localities in central western Queensland,
representing two discrete regions. The first region is in the
Carnarvon Gorge/Bigge Range area, and the second region is in
the vicinity of the Blackdown Tablelands. It is currently
unknown whether the distribution of the species is continuous,
or whether these discrete regions represent disjunct populations,
because the number of specimens known for this new species
are currently limited. Our genetic data suggest that the two
FIG. 3. Bayesian consensus phylogram of nuclear DNA (RAG1 locus) clades within Diporiphora nobbi and a newly identified species (D. sp. nov.),
including estimated dates for specific nodes and a distribution map of each major clade. Nodal support is shown in the form of Bayesian posterior
probabilities from 4 million generations, and estimated nodal dates are median values with 95%confidence intervals displayed in brackets from 10
million generations. This analysis confirms the mtDNA result that D. nobbi and D. sp. nov. are reciprocally monophyletic sister lineages.
SYSTEMATICS OF THE DIPORIPHORA NOBBI SPECIES COMPLEX 537
regions occupied by the species are genetically distinct;
however, this may simply be a function of isolation by distance
rather than suggestive of discrete genetic populations due to
limited sampling in intermittent areas. Genetic and morpholog-
ical analyses suggest that D. phaeospinosa is allopatric with
respect to the distribution of its congener D. nobbi. Further
analysis of the species distribution would be required to resolve
Etymology.—The specific epithet phaeospinosa is a composite
from the Latin root terms phaeo (dark) and spinos (spiny)
describing the main distinguishing morphological features of
Using an analysis of morphology and genetics, we show
extensive diversity within D. nobbi and uncover a previously
undescribed species within this species group, D. phaeospinosa.
This new species is highly divergent from nominal D. nobbi
individuals by using both molecular (Figs. 2, 3) and morpho-
logical characters (Fig. 4). We also describe high levels of
genetic diversity within nominal D. nobbi associated with
distinct geographic regions and habitat types across the broad
distribution of this species. Here, we describe the ramifications
of our work on perceived species boundaries within the D.
nobbi species complex, and we discuss possible hypotheses to
explain the potential diversification mechanisms within the D.
nobbi species complex and the biogeographic diversity within
nominal D. nobbi.
Species Boundaries and Taxonomy within the D. nobbi
Species Group.—Our analysis of molecular and morphological
variation within D. nobbi shows no support for current
subspecies delineations, D. n. nobbi and D. n. coggeri, within
the D. nobbi complex. Both holotypes for these subspecies seem
to occur within the geographic region encompassed by the
western NSW slopes and highland forests mtDNA subclade
(Fig. 2) and also cannot be resolved as subspecies using the
RAG1 dataset (Fig. 3). Similarly, there is no support for the
previously described W. parnabyi (Wells and Wellington, 1985),
later synonymized within D. nobbi, as distinct from D. nobbi
(Shea and Sadlier, 1999). This holotype is probably associated
with the mtDNA subclade from central western QLD. Our
results show that there is considerable diversity within
nominate D. nobbi. However, with the current data, an analysis
of morphological boundaries within the D. nobbi lineage and
among molecular subclades was not possible due to low
number of specimens with material available for both
molecular and morphological analyses.
Even with an analysis of morphological diversity within
nominate D. nobbi, it is unlikely that the currently described
subspecies, D. n. nobbi and D. n. coggeri (Witten, 1972) would be
resolved as morphologically distinct species or in fact distinct
subspecies, and we recommend that the two subspecies be
synonymized under the name of D. nobbi. Alternatively,
differences between these two groups may be a function of
genetic subdivision within this region observed within our
own mtDNA data (Fig. 2) and also observed within many other
species throughout the western NSW slopes region (Chapple et
al., 2005; O’Meally and Colgan, 2005; Colgan et al., 2009). Given
the high levels of molecular diversity within nominate D. nobbi,
TABLE 3. Summary of principal component (PC) loadings across
morphological variables for both male and female Diporiphora nobbi and
D. sp. nov. analyses, respectively. Values in bold indicate
measurements that are considered important (PC loading ,0.5).
Variable Male PC1 Male PC2 Female PC1 Female PC2
NS 3.749 21.241 2.9 0.345
RFP 1.131 0.533 0.854 20.238
LFP 1.011 0.301 0.704 20.238
RPAP 0.592 0.192 0.302 20.332
LPAP 0.647 0.275 0.393 20.199
MCS 20.179 21.08 20.13 1.179
MTS 20.691 22.083 20.311 1.632
ILS 0.452 20.163 — —
SLS 0.381 20.202 0.021 0.543
PAS 0.762 0.373 0.483 20.264
UAS 0.86 0.073 0.689 0.742
SUAS 0.855 0.231 — —
SS 0.771 0.806 0.515 20.544
NSP 1.002 0.255 0.987 0.07
PVS — — 0.171 0.065
RPVS — — 0.182 0.023
SBWPVS 0.589 0.297 0.569 0.018
TC 1.148 0.327 — —
Eigenvalue 23.3 8.71 12.33 5.66
explained 47.22 17.65 40.59 18.63
FIG. 4. Principal components analysis of male (a) and female (b)
morphological data based on genetic species described in the genetic
analyses. Principal components (PCs) are calculated from a covariance
matrix on an optimal set of morphological characters. PC1 is plotted
against PC2 in Euclidian space. PC1 explains 47.22 and 40.59%of the
variation and values are significantly different between D. nobbi and D.
sp. nov. in the male and female data, respectively. PC2 explains 17.65
and 18.63%of the variation in the male and female data, respectively.
PC2 is only significantly different between D. nobbi and D. sp. nov. for
the male data. Refer to Table 3 for an explanation of the variable
loadings for PC1 and PC2 for both the male and female data.
538 D. L. EDWARDS AND J. MELVILLE
it is possible that future molecular and morphological analyses
could find support for raising D. parnabyi and several other
subclades within nominate D. nobbi to species status. However,
there is no support warranting the use of a distinct genus
name, as ascribed by Wells and Wellington (1985).
Diversification and Biogeography of the D. nobbi Species
Complex.—Our divergence analyses suggest that the split
between D. nobbi and D. phaeospinosa occurred between 4 and
14 mya, with splits between the D. nobbi lineage and the D.
australis/bilineata lineage occurring at a similar time (between 8
and 19 mya). Due to the similar timing of these events, it is
likely that the developing aridity in Australia during the mid-
late Miocene, leading to dramatic changes in forest types
(Bowler, 1976; Macphail, 1997; Kershaw et al., 2003), is
associated with diversification of the eastern Australian
Diporiphora australis-nobbi lineage. Previous analysis has sug-
gested a northern origin for the D. australis lineage (Edwards
and Melville, 2010), which has only recently been present in
FIG. 5. Diagnostic drawings of morphological features distinguishing between Diporiphora nobbi (a, c, e, g, i) and Diporiphora phaeospinosa (b, d, f,
h, j). Illustrations show the arrangement of postauricular spines, lateral view, stylized (a–b); upper auricular spines and nuchal spine arrangement,
dorsal view, detailed (c–d), paravertebral and dorso-lateral scale keeling and regularity, dorsal view, stylized (e–f); dorso-lateral scale keeling, lateral
view, detailed (g–h); and differences between strength of scale keeling, close-up (i–j). Illustrations by Corrine Edwards.
SYSTEMATICS OF THE DIPORIPHORA NOBBI SPECIES COMPLEX 539
southern QLD and northern NSW as of the mid-late Pleistocene
(Hocknull et al., 2007). Given the much deeper age of nodes
within the D. nobbi species complex (Figs. 2, 3) this lineage
probably occupied a more southerly distribution historically,
and these two sister lineages have only recently been almost
entirely sympatrically distributed.
Both D. australis and D. nobbi currently occupy dry woodland
habitats, habitats that rapidly expanded during the mid-late
Miocene climate shift (Bowler, 1976; Macphail, 1997; Kershaw et
al., 2003). Diporiphora phaeospinosa, in contrast, seems to be
restricted to the wetter eucalypt forests in central southern QLD,
a region harboring many endemic and relictual species of snail
(A. F. Hugall, pers. comm.). Diversification within the D. nobbi
species complex is probably a result of sweeping climatic and
environmental change across the Australian continent at this
time (Bowler, 1976; Macphail, 1997; Kershaw et al., 2003) that
either led to vicariance between the D. nobbi and D. phaeospinosa
populations or to adaptive evolution within D. nobbi to favor the
broadly distributed dry woodland habitats coincident with
expansion into these newly available habitats. Many groups of
Australian lizards have shown rapid adaptive radiations during
the mid-late Miocene period, including other members of the
subfamily Amphibolurinae (e.g., Ctenophorus, Melville et al.,
2001), Elapidae (Sanders et al., 2008), and much more diverse
genera, such as Lerista (Skinner and Lee, 2009) and Ctenotus
(Rabosky et al., 2007).
Unlike D. australis (Edwards and Melville, 2010) and some
other northern QLD woodland taxa, the distribution of genetic
variation within D. nobbi cannot be predicted by known
biogeographic barriers. This is consistent with the hypothesis
that biogeographic patterns in woodland taxa may be harder to
predict than those of rainforest taxa (James and Moritz, 2000).
Rather, our mtDNA genetic data suggest that diversity within D.
nobbi is associated with specific habitat types. mtDNA clades
within D. nobbi are specifically associated with the northern
QLD Tablelands forests, central western QLD woodlands,
coastal forests and swamps of southeastern QLD and northeast-
ern NSW, western slopes and highland forests of NSW, central
NSW woodlands and the Murray River basin mallee–spinifex
woodlands of western NSW, eastern South Australia (SA), and
northwestern Victoria (VIC). Whereas our current nuclear data
do not show a similar pattern, the marker used here is not
particularly appropriate for testing population level associa-
tions. Our results provide preliminary evidence that mtDNA
divergence among D. nobbi subclades may be associated with
ecological differentiation, a response common in many of the
adaptive radiations reported from rapidly evolving arid regions
in Australia (Melville et al., 2001, 2006; Byrne et al., 2008).
However, to test this hypothesis increased genetic and morpho-
logical sampling in conjunction with ecological information is
required from across the distribution of the species.
Previous population genetic studies using both mtDNA and
allozyme markers have shown deeply divergent breaks among
D. nobbi populations in one region of NSW, which was also
identified in the current study. Driscoll and Hardy (2005)
alluded to a distinction between D. nobbi populations occupy-
ing a fragmented agricultural landscape versus continuous
mallee–spinifex woodlands in southern central NSW and
separated by the Lachlan River, in a region associated with
meeting range limits of many sister taxa without any obvious
steep ecological gradient. The two divergent populations were
not exchanging genes, and both were occupying mallee
woodlands with a spinifex understorey. Driscoll and Hardy
(2005) probably identified the junction between what we have
termed the central western NSW woodland clade and the
mallee–spinifex clade; however, both are occupying the same
mallee–spinifex habitat. Driscoll and Hardy (2005) also
uncovered weak evidence for increased dispersal within the
agricultural landscape. It is possible that western woodland
populations may have been able to move into the mallee
habitat postdisturbance and local extinction of the woodland
clade; alternatively, the Lachlan River may be providing the
divergence mechanism between these two clades as opposed to
simply habitat differentiation.
Our study also identifies high levels of differentiation among
D. nobbi populations across QLD and NSW, with mtDNA
subclades associated with coastal forests and swamps of
southeastern QLD/northeastern NSW, northern QLD wet/
dry tropical areas, central QLD woodlands, and the western
slopes of NSW. Divergence among the former two subclades
could be associated with the St. Lawrence Gap, seen in other
woodland (James and Moritz, 2000; Edwards and Melville,
2010) and rain forest taxa (Stuart-Fox et al., 2001; Moussalli et
al., 2005; Joseph and Omland, 2009). However, divergences
across this barrier are generally considered to correspond to the
Pleistocene (James and Moritz, 2000; Stuart-Fox et al., 2001;
Moussalli et al., 2005; Joseph and Omland, 2009; Edwards and
Melville, 2010). Alternatively, divergence among these QLD
and NSW subclades could have arisen through differentiation
across steep environmental and habitat gradients.
A combination of morphological and molecular information
is not only invaluable in the description of cryptic diversity but
also provides insight into the evolutionary processes driving
diversification within closely related species. Alpha and higher
level taxonomy in morphologically homoplasic groups, e.g.,
Amphibolurinae, require such information for accurate taxo-
nomic revision. The taxonomic history of the D. nobbi species
complex and cryptic diversity habored within it is indicative of
this problem in amphibolurine alpha taxonomy. Although our
study identifies some potential hypotheses explaining diversi-
fication within the D. nobbi species complex and population
differentiation within D. nobbi, this study is by no means a
complete treatment of diversity within this lineage. Much more
information is required to definitively test diversification
hypotheses within D. nobbi. This is true for eastern Australian
woodland environments in general that contain many widely
distributed species but that may, in fact, be species complexes,
crossing steep environmental gradients.
Acknowledgments.—We thank Steve Donnellan (South Aus-
tralian Museum), Andrew Amey, Patrick Couper (both of
Queensland Museum), and Ross Sadlier (Australian Museum)
for providing access to tissues and specimens. DE thanks Mark
Hutchinson (South Australian Museum) for discussions re-
garding morphology, Rebecca Rose and Katie Smith (both of
Museum Victoria) for assistance with field collections, Corrine
Edwards for illustrations, and Kate Sanders and Andrew
Hugall (both of University of Adelaide) for analytical discus-
sions. This research was funded by an Australian Research
Council Discovery Grant and an Australian Biological Re-
sources Study grant (to JM). Field collection and techniques
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Committee and Queensland Museum Animal Ethics Commit-
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APPENDIX 1. Genetic samples used in molecular analyses. Haplotype numbers, GenBank numbers, registration details, and site details are
outlined for each sample. Samples also used in morphological analyses are specified.
Lab no. Haplotype
no. Tissue no. Museum Latitude Longitude
DLE129 N1 JN627101 — N — 25 km N Alpha — Z10040 Mus. VIC 223.4822 146.6480
DLE135 N2 JN627050 — Y F 25 km N Alpha D74123 Z10044 Mus. VIC 223.4822 146.6480
DLE136 N1 JN627052 JN627123 Y M 25 km N Alpha D74124 Z10045 Mus. VIC 223.4822 146.6480
DLE137 N1 JN627085 JN627113 Y M 25 km N Alpha D74125 Z10046 Mus. VIC 223.4807 146.6452
DLE140 P1 JN627087 — Y M Bauhinia Stn. D74128 Z10049 Mus. VIC 225.1743 149.2046
DLE141 P2 JN627088 JN627115 Y M Bauhinia Stn. D74129 Z10050 Mus. VIC 225.1865 149.1618
DLE145 N3 JN627110 JN627117 Y M Maryborough D74132 Z10054 Mus. VIC 225.6046 152.8121
DLE146 N3 JN627111 JN627125 Y M Maryborough D74133 Z10055 Mus. VIC 225.6046 152.8121
DLE154 N4 JN627105 JN627127 N — Pilliga Forest, Narribri — Z10063 Mus. VIC 230.5657 149.4522
DLE161 N5 JN627086 JN627128 Y M Pilliga Forest, Narribri D74139 Z10070 Mus. VIC 230.5457 149.5389
DLE162 N6 JN627089 — Y M Pilliga Forest, Narribri D74140 Z10071 Mus. VIC 230.5667 149.4513
DLE164 N7 JN627094 — N — 140 km N Ivanhoe — Z10073 Mus. VIC 231.8998 143.5610
DLE167 N8 JN627045 — N — Gol Gol — Z10185 Mus. VIC 233.3795 143.5503
DLE168 N7 JN627053 — Y M 140 km N Ivanhoe D74141 Z10186 Mus. VIC 231.8953 143.5599
DLE169 N9 JN627062 — Y M 140 km N Ivanhoe D74142 Z10187 Mus. VIC 231.8999 143.5607
DLE170 N8 JN627091 — Y M Gol Gol D74143 Z10188 Mus. VIC 233.3796 143.5502
DLE171 N8 JN627063 JN627119 Y F Gol Gol D74144 Z10189 Mus. VIC 233.3796 143.5502
DLE172 N10 JN635331 JN627124 Y F West Wyalong D74145 Z10190 Mus. VIC 233.9821 147.0816
DLE173 N10 JN627108 — Y F West Wyalong D74146 Z10191 Mus. VIC 233.9823 147.0833
DLE174 N11 JN627081 — Y F Bolivia Hill R 134983 269 Aust. Mus. 229.4833 151.9000
DLE175 N12 JN627028 — Y M 19.7 km N Coombah
R 130998 338 Aust. Mus. 232.8167 141.6167
DLE176 N13 JN627054 — Y F 19.7 km N Coombah
R 130999 339 Aust. Mus. 232.8167 141.6167
DLE177 N14 JN627093 JN627121 N J 35 km W Yass R 135336 548 Aust. Mus. 234.9500 148.5333
DLE178 N15 JN627046 JN627120 Y F 90 km NE Bourke R 141023 1283 Aust. Mus. 229.3714 146.2333
DLE179 N16 JN627064 — N J 90 km NE Bourke R 141024 1284 Aust. Mus. 229.3431 146.2314
DLE180 N17 JN627065 — Y M 90 km NE Bourke R 141025 1285 Aust. Mus. 229.3611 146.2242
DLE181 N18 JN627034 — N J 90 km NE Bourke R 141026 1286 Aust. Mus. 229.3833 146.1500
DLE182 P3 JN627029 JN627116 Y F Blackdown Tbld. R 151842 5009 Aust. Mus. 223.7911 149.0939
DLE183 P3 JN627041 — Y M Blackdown Tbld. R 151843 5010 Aust. Mus. 223.7586 149.1025
DLE184 P3 JN627038 JN627114 Y M Blackdown Tbld. R 151844 5011 Aust. Mus. 223.7586 149.1025
DLE185 P3 JN627030 JN627133 Y F Blackdown Tbld. R 151845 5012 Aust. Mus. 223.7586 149.1025
DLE186 N19 JN627109 — Y M Cassilis-Coolah Rd. R 152062 5301 Aust. Mus. 232.0550 149.9333
DLE187 N20 JN627035 — Y M Moonbi Ranges R 152046 5338 Aust. Mus. 230.9925 151.0833
DLE188 N21 JN627066 — N J Moonbi Ranges R 152079 5340 Aust. Mus. 230.9925 151.0833
DLE189 N21 JN627033 — N J Moonbi Ranges R 152080 5342 Aust. Mus. 230.9925 151.0833
DLE190 N22 JN627067 JN627134 Y M Torrington SF R 152098 5459 Aust. Mus. 229.3392 151.6914
DLE191 N23 JN627082 JN627112 N J Boonoo Boonoo NP R 152341 6129 Aust. Mus. 231.5564 152.1269
DLE193 N24 JN627047 JN627135 N J Pyes Creek Rd. R 159836 6488 Aust. Mus. 229.2353 151.8372
DLE194 N25 JN635332 — N J Bolivia Hill R 159768 6555 Aust. Mus. 229.3214 151.9181
DLE196 N26 JN627068 — Y M Warrakoo Stn. R 153217 7113 Aust. Mus. 233.9858 141.1181
DLE197 N27 JN627095 JN627130 Y F 2 km N Tamworth R 157284 7232 Aust. Mus. 231.0850 150.9472
DLE198 N28 JN627042 — Y F 2 km N Tamworth R 157285 7233 Aust. Mus. 231.0850 150.9472
DLE199 N29 JN627037 — Y F 2 km N Tamworth R 157286 7234 Aust. Mus. 231.0850 150.9472
DLE200 N20 JN627097 — Y F 2 km N Tamworth R 157287 7235 Aust. Mus. 231.0850 150.9472
DLE201 N30 JN627036 — Y F 2 km N Tamworth R 157288 7236 Aust. Mus. 231.0850 150.9472
DLE202 N31 JN627106 — Y M Warrumbungle NP R 156048 7613 Aust. Mus. 231.3289 148.9967
DLE203 N32 JN627055 — N J Torrington SF R 157100 7933 Aust. Mus. 229.2944 151.6772
DLE204 N33 JN627056 — Y M Torrington SF R 157201 8121 Aust. Mus. 229.2944 151.6772
DLE205 N34 JN627102 — Y M Yarrowick R 157018 8150 Aust. Mus. 230.4725 151.3714
DLE206 N35 JN627031 — Y M Yarrowick R 157023 8154 Aust. Mus. 230.4725 151.3714
DLE207 N35 JN627103 — Y M Yarrowick R 157024 8156 Aust. Mus. 230.4725 151.3714
DLE208 N36 JN627032 — Y F Bundarra-Inverell Rd. R 157074 8161 Aust. Mus. 230.0747 151.0967
DLE209 N37 JN627099 JN627131 Y M Barraba Rd. R 157216 8162 Aust. Mus. 230.1422 150.7872
DLE210 N37 JN627069 JN627129 Y M Barraba Rd. R 157217 8164 Aust. Mus. 230.1450 150.7869
DLE211 N24 JN627048 — N J Bolivia Hill R 157221 8170 Aust. Mus. 229.3372 151.8944
DLE212 N38 JN627070 — N J Bolivia Hill R 157232 8192 Aust. Mus. 229.3264 151.9631
DLE213 N39 JN627027 — Y M Copeton R 157218 8307 Aust. Mus. 229.9167 151.0167
DLE214 N40 JN627071 — N — 35 km from Mt Hope R 156632 8732 Aust. Mus. 232.9467 146.1922
DLE215 N41 JN627025 — Y F 35 km from Mt Hope R 156683 8777 Aust. Mus. 232.8622 146.1894
DLE216 N41 JN627057 JN627132 Y F 35 km from Mt Hope R 156684 8778 Aust. Mus. 232.8622 146.1894
DLE217 N41 JN627058 — Y M 35 km from Mt Hope R 156685 8779 Aust. Mus. 232.8622 146.1894
542 D. L. EDWARDS AND J. MELVILLE
Lab no. Haplotype
no. Tissue no. Museum Latitude Longitude
DLE218 N40 JN627051 — Y F 35 km from Mt Hope R 156686 8780 Aust. Mus. 232.8622 146.1894
DLE231 N42 JN627107 — N J Gundabooka NP R 164801 — Aust. Mus. 230.5000 145.7167
DLE232 N43 JN627083 JN627122 Y M Airlie Beach R 164849 — Aust. Mus. 220.2667 148.7000
DLE242 N44 JN627092 — Y F Kirrama J82745 — QLD Mus. 218.1536 145.6833
JEM196 N12 JN627104 — Y F Wyperfeld NP D71317 Z10192 Mus. VIC 235.4594 142.0069
JEM197 N45 JN627078 — N — Wyperfeld NP D71318 Z10193 Mus. VIC 235.4594 142.0069
JEM198 N12 JN627084 — N — Wyperfeld NP D71319 Z10194 Mus. VIC 235.4594 142.0069
JEM199 N46 JN627079 — N — Wyperfeld NP D71320 Z10195 Mus. VIC 235.4594 142.0069
JEM200 N12 JN627061 — N — Wyperfeld NP D71321 Z7876 Mus. VIC 235.4594 142.0069
JEM222 N47 JN627090 — N — Murray-Sunset NP D71343 Z7875 Mus. VIC 234.7597 141.7758
JEM223 N12 JN627075 — N — Murray-Sunset NP D71344 Z7874 Mus. VIC 234.7597 141.7758
JEM224 N12 JN627076 — N — Murray-Sunset NP D71345 Z7873 Mus. VIC 234.7597 141.7758
JEM225 N48 JN627080 — N — Murray-Sunset NP D71346 Z7872 Mus. VIC 234.7597 141.7758
JEM226 N12 JN627077 — N — Murray-Sunset NP D71347 Z7871 Mus. VIC 234.7597 141.7758
JEM299 N45 JN627044 — N — Hattah NP D71420 Z7870 Mus. VIC 234.6853 142.2797
JEM300 N49 JN627100 JN627118 N — Hattah NP D71421 Z7868 Mus. VIC 234.6853 142.2797
JEM301 N50 JN627098 — N — Hattah NP D71422 Z7867 Mus. VIC 234.6853 142.2797
JEM302 N51 JN627096 — N — Hattah NP D71423 Z7869 Mus. VIC 234.6853 142.2797
DLE243 N52 JN627039 JN627126 N — Rainbow Beach — ABTC03912 SA Mus. 225.9060 153.0790
DLE244 N53 JN627059 — N — Rainbow Beach — ABTC03913 SA Mus. 225.9060 153.0790
DLE245 N53 JN627049 — N — Rainbow Beach — ABTC03914 SA Mus. 225.9060 153.0790
DLE246 N53 JN635333 — N — Rainbow Beach — ABTC03915 SA Mus. 225.9060 153.0790
DLE247 N54 JN627026 — Y F Kroombit Tops J54855 ABTC24230 QLD Mus. 224.3667 151.0333
DLE248 N12 JN627060 — Y M N Mulga Stn. R22791 ABTC53153 SA Mus. 230.2000 139.7000
DLE249 N55 JN627072 — Y M N Mulga Stn. R22792 ABTC53154 SA Mus. 230.2000 139.7000
DLE250 N56 JN627043 — N J Swan Reach CP R26165 ABTC53191 SA Mus. 234.6000 139.4833
DLE251 N57 JN627040 — Y M Swan Reach CP R26902 ABTC53201 SA Mus. 234.6000 139.4833
DLE252 N12 JN627073 — Y M Arkaroola R52947 ABTC74088 SA Mus. 230.1206 139.3986
DLE253 N12 JN627074 — Y F Arkaroola R52940 ABTC74147 SA Mus. 230.1206 139.3986
Y, yes; N, no.
M, male; F, female; J, juvenile.
Mt, mountains; Rd., Road, Stn., Station; Tbld., Tablelands.
APPENDIX 1. Continued.
SYSTEMATICS OF THE DIPORIPHORA NOBBI SPECIES COMPLEX 543
APPENDIX 2. Registration, sex, species, and locality information for all samples used only for morphological analyses. Coordinates are given in
Latitude Longitude Museum
2636 F D. nobbi Crows Nest 227.2667 152.0500 Aus. Mus.
11675 F D. nobbi Kainkillenbun 227.1000 151.5333 Aus. Mus.
16955 M D. nobbi Cooranga north 224.7667 151.4000 Aus. Mus.
16959 M D. nobbi Bunya Mountains 226.8500 151.5667 Aus. Mus.
30358 F D. nobbi Hillston 233.4833 145.5333 Aus. Mus.
65949 M D. nobbi 88 km W Alpha 223.5667 145.7833 Aus. Mus.
73578 F D. nobbi Bathurst 233.4167 149.5833 Aus. Mus.
74767 M D. nobbi 4 m S Stanthorpe 228.7333 141.8667 Aus. Mus.
74768 M D. nobbi 4 m S Stanthorpe 228.7333 141.8667 Aus. Mus.
76492 M D. nobbi 27 km N Albury 235.9667 147.0167 Aus. Mus.
80246 F D. nobbi 15 km W Baldry 232.8167 148.3667 Aus. Mus.
81686 F D. nobbi Monoboli NR 232.2667 150.8833 Aus. Mus.
116024 F D. nobbi Melbergen Range 233.8500 146.0333 Aus. Mus.
130061 M D. nobbi 26.5 km N Cudgegong River 232.5667 150.0833 Aus. Mus.
141023 F D. nobbi Beulah Stn. 229.3714 146.2333 Aus. Mus.
141025 M D. nobbi Beulah Stn. 229.3611 146.2242 Aus. Mus.
141585 F D. nobbi Kangaroo River SF 230.1167 152.8333 Aus. Mus.
142078 F D. nobbi Beaury SF 228.5469 152.3364 Aus. Mus.
158578 F D. nobbi —230.9544 149.5350 Aus. Mus.
161978 F D. nobbi Mt Kaputar NP 230.2375 150.0897 Aus. Mus.
162105 F D. nobbi Playgan SF 230.4464 150.2817 Aus. Mus.
941 F D. nobbi Goombungee 227.5333 151.2833 QLD Mus.
1083 M D. nobbi Batavia River 212.1833 141.9000 QLD Mus.
1084 M D. nobbi Batavia River 212.1833 141.9000 QLD Mus.
10493 M D. nobbi Inverell 229.7667 151.1167 QLD Mus.
10494 M D. nobbi Inverell 229.7667 151.1167 QLD Mus.
30734 F D. nobbi Texas Caves 228.8833 151.4333 QLD Mus.
32346 F D. nobbi Boonoo Boonoo Falls 228.8000 152.1667 QLD Mus.
35150 M D. nobbi Airlie Beach 220.2667 148.7167 QLD Mus.
36887 F D. nobbi Epping Forest 222.3167 146.7500 QLD Mus.
36893 M D. nobbi Nine Mile 223.8333 147.4500 QLD Mus.
37117 F D. nobbi Springsure 224.1333 147.9167 QLD Mus.
38749 M D. nobbi Mt Windsor Tbld. 216.3167 145.0167 QLD Mus.
40123 M D. nobbi Kroombit Tops 224.3667 150.9833 QLD Mus.
42144 M D. nobbi Kroombit Tops 224.3667 150.9833 QLD Mus.
42178 M D. nobbi Kroombit Tops 224.3667 150.9833 QLD Mus.
44358 F D. nobbi Victoria Downs North 220.7167 146.4500 QLD Mus.
44905 M D. nobbi Helenslee Stn. 220.5167 145.7000 QLD Mus.
46702 M D. nobbi Myross Stn., 25 km N Aramac 222.8167 145.3667 QLD Mus.
46723 F D. nobbi Winhaven Stn. 222.9500 145.6833 QLD Mus.
47798 F D. nobbi 10 km S Barcaldine 223.6167 145.2833 QLD Mus.
53009 M D. nobbi Mt Cleveland 219.2500 147.0167 QLD Mus.
56060 M D. nobbi Ka Ka Mundi NP 224.8167 147.4000 QLD Mus.
62953 M D. nobbi The Bluff, Keysland 226.2333 151.7000 QLD Mus.
63109 M D. nobbi Blair Athol Coal Mine 222.7000 147.5500 QLD Mus.
64762 F D. nobbi Charters Towers 220.0833 146.2667 QLD Mus.
74745 M D. nobbi Bauple SF 225.8167 152.6167 QLD Mus.
74746 M D. nobbi Cunningham’s Gap NP, Mt
228.0333 152.3833 QLD Mus.
75454 M D. nobbi Kirrama 218.1500 145.6167 QLD Mus.
76743 F D. nobbi Cudmore NP 222.9333 146.3667 QLD Mus.
62741 M D. nobbi Mt Abbot 220.1000 147.7500 QLD Mus.
77780 M D. nobbi Southward NP 227.8300 150.1000 QLD Mus.
32596 M D. phaeospinosa Blackdown Tbld. 223.8000 149.1333 QLD Mus.
33335 M D. phaeospinosa Blackdown Tbld. 223.8000 149.0667 QLD Mus.
33336 M D. phaeospinosa Blackdown Tbld. 223.8000 149.0667 QLD Mus.
34294 F D. phaeospinosa Blackdown Tbld. 223.8000 149.1333 QLD Mus.
34295 F D. phaeospinosa Blackdown Tbld. 223.8000 149.1333 QLD Mus.
34296 M D. phaeospinosa Blackdown Tbld. 223.8000 149.1333 QLD Mus.
36891 M D. phaeospinosa Blackdown Tbld. 223.8000 149.1333 QLD Mus.
36892 F D. phaeospinosa Reklau Pk. 223.3333 147.5000 QLD Mus.
38591 F D. phaeospinosa Glenhaughton Stn 225.2333 148.9500 QLD Mus.
50807 F D. phaeospinosa Blackdown Tbld. 223.8000 149.1000 QLD Mus.
28495 J D. phaeospinosa Blackdown Tbld. 223.8000 149.1333 QLD Mus.
30267 J D. phaeospinosa Robinson Gorge 225.2833 149.1500 QLD Mus.
38560 J D. phaeospinosa Robinson Gorge 225.2833 149.1500 QLD Mus.
38590 J D. phaeospinosa Robinson Gorge 225.2833 149.1500 QLD Mus.
36890 F D. phaeospinosa Blackdown Tbld. 223.8000 149.1333 QLD Mus.
38589 J D. phaeospinosa Glenhaughton Stn. 225.2333 148.9500 QLD Mus.
M, male; F, female; J, juvenile.
Pk., peak; Mt, mountains; NP, National Park; Rd., Road, Stn., Station; Tbld., Tablelands.
544 D. L. EDWARDS AND J. MELVILLE
APPENDIX 3. Ultrametric BEAST output tree for the RAG1 analyses. GenBank numbers for sequences are provided. Node ages are shown with bars representing 95%confidence intervals of node
SYSTEMATICS OF THE DIPORIPHORA NOBBI SPECIES COMPLEX 545
APPENDIX 4. Ultrametric BEAST output for mtDNA analyses. GenBank numbers or registration numbers of specimens are shown. See Appendix 1 for corresponding GenBank numbers for these
specimens. Node ages also are shown, with 95%HPD errors around node ages being shaded.
546 D. L. EDWARDS AND J. MELVILLE