2 EVOLUTION OF CAVIOMORPH RODENTS:
A COMPLETE PHYLOGENY AND TIMETREE
FOR LIVING GENERA
EVOLUCIÓN DE LOS ROEDORES CAVIOMORFOS: FILOGENIA Y
ÁRBOL TEMPORAL COMPLETO DE LOS GÉNEROS VIVIENTES
Nathan S. UPHAM1, 2, 3 and Bruce D. PATTERSON2
1Committee on Evolutionary Biology, University of Chicago, IL 60637 Chicago, USA. email@example.com
2Integrative Research Center, Field Museum of Natural History, IL 60605 Chicago, USA. firstname.lastname@example.org
3Ecology and Evolutionary Biology, Yale University, CT 06511 New Haven, USA.
Abstract. e Caviomorpha is a diverse lineage of hystricognath rodents endemic to the Americas and
Caribbean islands. We analyzed evolutionary relationships within 11 families of caviomorphs and their
relatives in the suborder Ctenohystrica using a supermatrix of 199 taxa and DNA sequences from ve
genes. New gene sequences were generated for 33 genera, including 12 genera newly available for molecular
analysis. Presented here are the analyses pruned to a single representative for each genus, totaling 68 of the
70 living genera in Ctenohystrica. Our analyses recovered strong support for Hystricognathi containing
the monophyletic groups Hystricidae, Phiomorpha, and Caviomorpha, with the latter two groups as well-
supported sister taxa. e analyses also strongly supported the monophyly of the four traditional superfamilies
of caviomorphs, with Cavioidea + Erethizontoidea and Chinchilloidea (including Dinomyidae) +
Octodontoidea. Cuniculidae + Dasyproctidae are recovered as sister to Caviidae (including Hydrochoerus).
Abrocomidae (including Cuscomys) is sister to the remaining octodontoid families, consisting of the
dyads Octodontidae + Ctenomyidae and Echimyidae (including Myocastor) + Capromyidae. e ve
genera of capromyids form a robustly monophyletic group, but they are allied to a group of Brazilian
echimyids, rendering Echimyidae paraphyletic. We dated nodes in our tree by comparing eight sets of
fossil calibrations, identifying a set of 22 calibrations that minimized internal age conicts.e resulting
timetree dates the Hystricognathi crown to the Middle Eocene, 44.9 Ma, and the phiomorph-caviomorph
split to 42.0 Ma. Crown caviomorphs diverged at 35.7 Ma, and splits of Cavioidea-Erethizontoidea and
Chinchilloidea-Octodontoidea occurred at 32.4 Ma and 32.8 Ma, respectively. Most families appeared
in the late Oligocene-Early Miocene and virtually all genera are of Middle-Late Miocene age, with a
few exceptions. We briey consider geo-climatic changes that might have inuenced the evolution of
hystricognath rodents, deferring to another work a detailed analysis of their rates and ecological drivers
Resumen. Los caviomorfos constituyen un linaje diverso de roedores histricognatos endémicos de las
Américas y las islas del Caribe. En este estudio, analizamos secuencias de ADN (mitocondrial y exones
nucleares) de cinco genes en el suborden Ctenohystrica, incluyendo roedores de las familias Diatomyidae,
Ctenodactylidae, Hystricidae, Petromuridae, ryonomyidae, Bathyergidae, y 11 familias de caviomorfos.
Se generaron nuevas secuencias de cyt-b, 12S rRNA, GHR, vWF, y/o RAG1 para 33 géneros, 31 de estos
caviomorfos, 12 de las cuales corresponden a secuencias nuevas disponibles para análisis logenéticos. Se
utilizó una supermatriz de 199 taxa y 5194 pb de ADN para el análisis de las relaciones evolutivas del
SAREM Series A - Mammalogical Research, Vol 1 2015 63
Upham and Patterson
grupo. En este estudio, se redujeron los análisis a un solo representante por cada género, incluyendo un
total de 68 de los 70 géneros vivientes de Ctenohystrica. Nuestros análisis recuperan un fuerte soporte
para Hystricognathi: puercoespines del Viejo Mundo + omorfos + caviomorfos, con monolia para los
tres grupos: Phiomorpha con Bathyergidae y hermano de Petromuridae + ryonomyidae; y los omorfos
como grupo hermano de Caviomorpha. Los análisis también soportan fuertemente la monolia de las
cuatro superfamilias tradicionales de caviomorfos, con Cavioidea como hermana de Erethizontoidea y
Chinchilloidea (incluyendo Dinomyidae) como hermana de Octodontoidea. Cuniculidae + Dasyproctidae
fueron recuperadas como grupo hermano de Caviidae (incluyendo Hydrochoerus). Abrocomidae
(incluyendo Cuscomys) aparece como hermana del resto de las familias de octodontoideos, consistiendo en
las siguientes díadas: Octodontidae + Ctenomyidae y Echimyidae (incluyendo Myocastor) + Capromyidae.
Aunque cinco géneros de caprómidos conforman un robusto grupo monolético, fueron recuperados
como grupo hermano de los equímidos brasileros (Carterodon, Clyomys, Euryzygomatomys, y Trinomys)
quedando Echimyidae como paralético. Datamos los nodos de nuestro árbol, comparando ocho grupos
de calibraciones fósiles para identicar un grupo de 22 puntos de calibración, los que minimizaron los
conictos de edad entre las dataciones. El árbol calibrado resultante ubica el origen de la divergencia
del grupo corona de Hystricognathi en el Eoceno medio (44.9 Ma), poco antes de la separación de
Phiomorpha respecto de Caviomorpha hace 42.0 Ma. El grupo corona de caviomorfos divergió hace 35.7
Ma, separándose Cavioidea respecto de Erethizontoidea, y Chinchilloidea respecto de Octodontoidea hace
32.4 Ma y 32.8 Ma, respectivamente. El árbol datado muestra que la mayoría de las familias aparecieron en
el Oligoceno tardío o en el Mioceno temprano. Virtualmente todos los géneros tuvieron su origen hacia el
Mioceno medio a tardío, con algunas excepciones notables. Los antepasados de los caviomorfos habitaron
ambientes variados y dinámicos durante el Eoceno medio-Reciente, historia que probablemente conguró
su trayectoria evolutiva. La diversicación inicial de Caviomorpha en cuatro superfamilias fuertemente
diferenciadas coincide con un evento de enfriamiento global dramático, cerca del límite Eoceno-Oligoceno,
que representa la transición desde un “planeta invernadero” (caliente, húmedo y carente de casquetes
polares) a un planeta más frío y seco, con presencia de glaciaciones. Durante el Mioceno medio (~15
Ma), el levantamiento de los Andes Centrales creó un patrón de lluvias en gran parte del este de América
del Sur, con la transformación del continente, en términos generales, en biomas que fueron áridos en el
sur y mésicos en el norte, cada uno con desafíos ambientales distintivos para los caviomorfos residentes.
El registro fósil de Octodontoidea parece reejar estos cambios. Así, el clado Echimyidae-Capromyidae
adaptado a ambientes mésicos, se encuentra representado en las localidades de la Patagonia durante el
Mioceno, pero es reemplazado posteriormente por una mayor diversidad de formas adaptadas a ambientes
áridos, tales como Octodontidae-Ctenomyidae. Nuestro reloj molecular sugiere que estos clados principales
de Octodontoidea experimentaron diversicaciones que son aproximadamente coetáneas (18.2 Ma y 18.9
Ma, respectivamente), más o menos concordantes con el principio de la diferenciación climática en todo
el continente. Los datos espacio-temporales y ecomorfológicos del registro fósil necesitan correlacionarse,
adicionalmente, con la aridez progresiva de la región, sobre todo ahora que la secuencia de eventos de
diversicación del linaje se puede inferir a partir del reloj molecular. Nos resta abordar en futuros trabajos,
una evaluación más detallada del tiempo, el modo y los factores ecológicos que estuvieron asociados a la
diversicación biológica de los roedores caviomorfos.
Caviomorph evolution through time
Studying how and when lineages diversify is a central strategy for illuminating the his-
tory of life. Yet investigating the timing and drivers of diversication requires rst establishing
a common timescale of evolutionary and geologic events. Whereas molecular phylogenies yield
relative estimates of divergence timing in units of DNA substitutions, geologic ages (e.g., from
fossils) are needed to convert those molecular divergences to absolute time in years. Because the
oldest diagnostic fossil in a clade establishes its minimum age, molecular rates can be calibrated
to time and used with relaxed-clock models of among-species rate variation to estimate diver-
gence times (Drummond et al., 2006; Ho and Phillips, 2009). Phylogenetic trees that are scaled
to time (hereafter referred to as “timetrees;” Hedges and Kumar, 2009) allow comparisons of
timing and rates of diversication among clades, across taxa, and in relation to known geologic
events (e.g., Dávalos, 2010; Schenk et al., 2013).e ability to contrast patterns of evolution in
contemporaneous units enables the investigation of evolutionary tempo and mode from mo-
lecular phylogenies (Nee et al., 1992), and presents new ways to test long-standing hypotheses,
e.g., assesing the biogeographic origins of lineages (Ronquist, 1997; Ree et al., 2005).
Here we investigate the timescale of evolution across a speciose and fossil-rich clade of
Neotropical rodents (Hystricognathi: Caviomorpha). is American and Caribbean lineage is
an exceptionally diverse radiation comprised of four superfamilies, 11 families, and at least
244 living species. e Octodontoidea (spiny rats, degus, and their allies) are the most diverse,
comprising 70% of caviomorph genera and 75% of living species. However, Cavioidea (guinea
pigs and capybaras) and Chinchilloidea (chinchillas and pacaranas) both formerly exhibited
considerably greater diversity: each superfamily was represented by 20 genera during the Late
Miocene (11.2-5.3 Ma), alongside 18 genera of octodontoids from the same time period
(McKenna and Bell, 1997; Upham, 2014). Analyses of their diversity over time are, however,
beyond the scope of this paper and are instead the subject of a succeeding one. Here we focus
on the phylogenetic relationships of living caviomorph genera.
Caviomorph rodents and each of the four superfamilies that comprise this group are mainly
distributed in South America (Fig. 1). e overall pattern of caviomorph species richness (Fig.
1.1) is founded in that of Octodontoidea (Fig. 1.2); this group has concentrated richness in Ama-
zonia and the Atlantic Forest but is distributed from the Greater Antilles and Central America to
Tierra del Fuego and in most intervening habitats. Collectively, octodontoids exploit most niches
available to rodents, assuming tree-living (arboreal and scansorial), burrowing (semi-fossorial
and fossorial), terrestrial, rock-dwelling, and even semi-aquatic forms (Reig, 1981; Mares and
Ojeda, 1982). e next-most diverse lineage is the Cavioidea (Fig. 1.3), whose geographic range
is nearly as extensive as the octodontoids but with peak richness in the Guianan and Brazilian
highlands. Cavioids encompass remarkable morphological disparity (including near-cursorial
digitigrades and the largest living rodents), inhabit various ecosystems, and display diverse levels
of sociality (Rowe and Honeycutt, 2002; Perez and Pol, 2012). e Erethizontoidea (Fig. 1.4)
are widely distributed but only modestly diversied as medium-sized arboreal herbivores (Voss,
2011; Voss et al., 2013); this group includes the only caviomorph lineage to colonize Nearctic
portions of North America. Finally, the Chinchilloidea (Fig. 1.5) are represented by a couple of
distantly related lineages that inhabit arid habitats and wet forests, respectively, in western South
America (Mones, 1981; Spotorno et al., 2004).
Upham and Patterson
Figure 1. Species richness maps based on known geographic ranges and extant species of caviomorph rodents: 1.
Caviomorpha, and its four component superfamilies; 2. Octodontoidea; 3. Cavioidea; 4. Erethizontoidea; and 5. Chinchilloidea.
Species richness estimates are based on range overlap within 0.5 degree square grid cells, and are calculated using .shp les
updated from the IUCN (2012) database.
Some caviomorphs adopt lifestyles exploited elsewhere by distantly related rodent groups like
tree and ground squirrels, pocket gophers, and muskrats. Others may be ecological analogues
to hyraxes (Hyracoidea: Procaviidae), duikers (Bovidae: Cephalophinae), rabbits and hares
(Leporidae), hippos (Hippopotamidae), and chevrotains (Tragulidae; Mares and Ojeda, 1982).
All of these other groups were absent from South America during most of its Cenozoic isolation
as an island continent (Simpson, 1980). e sudden appearance of caviomorphs during the midst
of this isolation (Wyss et al., 1993; Campbell, 2004), their extensive fossil record back to the
Middle Eocene (McKenna and Bell, 1997; Antoine et al., 2012), and their modern ecological
diversity and morphological disparity make Caviomorpha an excellent lineage for investigating
biotic responses to changing landscapes (Veblen et al., 2007; Madden et al., 2010), climates
(Hooghiemstra and van der Hammen, 1998), and biotas (Goin et al., 2010).
We aim to construct a robust genus-level timetree from multiple genes and fossil calibra-
tions to enable assessment of the patterns, tempo, and modes of diversication of Caviomor-
pha. In the present chapter we seek to: 1) identify its higher-level relationships to sister lineages
in Africa and Asia; 2) identify and detail the relationships among all living caviomorph genera;
and 3) reconstruct the timing of diversication throughout the group’s Paleogene-to-Neogene
Caviomorph evolution through time
Materials and methods
e Suborder Ctenohystrica is one of three higher-level clades in Rodentia supported
by recent molecular phylogenetic studies, others being the “mouse-related” and “squirrel-
related” clades (Huchon et al., 2000; Huchon et al., 2007; Montgelard et al., 2008; Blanga-
Kan et al., 2009; Churakov et al., 2010; Fabre et al., 2012a; Wu et al., 2012). Some refer
to this group as Hystricomorpha (Carleton and Musser, 2005), but to dierentiate it from
the hystricomorphous condition of rodent masseter muscles (not shared by all members) we
employ the term Ctenohystrica (Huchon et al., 2000).e Ctenohystrica includes 18 of the
33 living families of rodents, and is divided into the Ctenodactylomorphi, consisting of two
small relict families in North Africa and Southeast Asia, and Hystricognathi, which consists of
16 tropical families, one reaching into North America (Carleton and Musser, 2005; Huchon
et al., 2007). Hystricognathi is then divided into the Old World porcupines (Hystricidae),
Phiomorpha in Africa (three living families of mole-rats, dassie rats, and cane rats), and
Caviomorpha in the Americas and Caribbean (11 living families, including Myocastor within
Echimyidae, and Hydrochoerus within Caviidae) (Huchon and Douzery, 2001; Carleton and
Musser, 2005; Martin, 2005; Opazo, 2005; Sallam et al., 2009; Fabre et al., 2012b; Upham
and Patterson, 2012; Honeycutt, 2013).
Here we present patterns of evolution among genera that are based upon phylogenetic
analyses conducted across 198 species in Ctenohystrica and presented in Upham (2014).
In this study, we describe all aspects of the phylogenetic reconstruction –including DNA
sequencing and alignments, GenBank accession numbers, gene combinability analyses, and
fossil calibrations– in order to facilitate future work on this fascinating group of mammals.
Our analyses included dense taxon sampling across all families in Ctenohystrica, employing
one representative per species and as many species as possible (Tab. 1; App. 1). We sampled 68
of 70 living genera in Ctenohystrica, all genera in Hystricognathi, and 69% of the living spe-
cies (Tab. 1). Although most species sampled were from Octodontoidea (126 species), species
sampling within superfamilies was roughly proportional to their extant diversity (62-88% of
species; Tab. 1). e springhaas Pedetes (Anomaluromorpha) was the outgroup for all phylo-
genetic analyses because it is a member of the mouse-related clade most likely to be sister to
Ctenohystrica (but see Huchon et al., 2007; Montgelard et al., 2008; Blanga-Kanet al., 2009;
Churakov et al., 2010; Fabre et al., 2012a). e classication, taxon and gene sampling, and
GenBank accession numbers for this study are presented in Appendix 1.
e gene sampling from two earlier studies of caviomorph rodent evolution (Upham and
Patterson, 2012; Upham et al., 2013) provided the foundation for this study. We generated an
assortment of new DNA sequences for ve genes that are listed in Appendix 1. ese include
two mitochondrial loci [mtDNA; cytochrome-b (cyt-b) and 12S ribosomal RNA (12S rRNA)]
and three unlinked nuclear exons [growth hormone receptor exon 10 (GHR), von Willebrand
factor exon 28 (vWF), and recombination activating gene 1 (RAG1)]. ese genes were selected
on the basis of variation in evolutionary rates (mitochondrial vs. nuclear), the diversity of taxa
Upham and Patterson
>ŝǀŝŶŐ ^ĂŵƉůĞĚ %>ŝǀŝŶŐ ^ĂŵƉůĞĚ %
dEK,z^dZ/ 70 68 97 288 198 69
CTENODACTYLOMORPHI 5 3 60 6 3 50
Ctenodactylidae 4 2 50 5 2 40
Diatomyidae 1 1 * 1 1 *
HYSTRICOGNATHI 65 65 * 282 195 69
Hystricidae 3 3 * 11 7 64
Bathyergidae 6 6 * 24 19 79
Petromuridae 1 1 * 1 1 *
Thryonomyidae 1 1 * 2 1 50
ò®ÊÃÊÙÖ« 54 54 * 244 167 68
Caviidae 6 6 * 19 14 74
Cuniculidae 1 1 * 2 2 *
Chinchillidae 3 3 * 7 5 71
Dinomyidae 1 1 * 1 1 *
KãÊÊÄãÊ® 38 38 * 186 126 68
Abrocomidae 2 2 * 9 4 44
Ctenomyidae 1 1 * 62 46 74
Echimyidae 22 22 * 89 56 63
Capromyidae 5 5 * 13 8 62
previously sampled, and their demonstrated utility in rodent phylogenetics (Huchon and
Douzery, 2001; Honeycutt et al., 2003; Galewski et al., 2005; Opazo, 2005; Patterson and
Velazco, 2008; Upham and Patterson, 2012). e use of cyt-b in this study lends breadth to our
taxon sampling, particularly at the species-level (e.g., in Ctenomyidae; Parada et al., 2011), that
has been lacking in evolutionary studies of Caviomorpha as a whole.
Table 1. Living diversity of rodent families in Ctenohystrica and the taxa sampled for one or more genes in this study. Taxonomy
and counts are updated from Woods and Kilpatrick (2005). An asterisk (*) signies all members sampled. Bold values dierentiate
higher clades from families.
Caviomorph evolution through time
DNA sequencing from fresh and dried tissue
We isolated genomic DNA from frozen fresh tissues (liver, kidney or muscle) preserved in ethanol,
or from dried tissues (muscle, skin) adhering to museum voucher specimens. Fresh tissue DNA was ex-
tracted from 37 specimens for this study, using either the DNeasy Blood & Tissue Kit (QIAGEN) or phe-
nol-chloroform methods, both following standard protocols. Molecular laboratory work on fresh tissues
was conducted in three labs depending on the source of the tissues: Pritzker Laboratory for Molecular
Systematics and Evolution (Field Museum of Natural History, Chicago, IL, USA), Laboratory of Molecular
Systematics (Royal Ontario Museum, Toronto, ON, Canada), and Laboratório de Mastozoologia e Bio-
geograﬁa (Universidade Federal do Espírito Santo, Vitória, ES, Brazil). Nucleic acid concentrations were
quantiﬁed using a NanoDrop spectrophotometer (Thermo Fisher Scientiﬁc).
Dried-tissue DNA, also called ancient DNA (aDNA), was extracted from eight specimens for this
study and analyzed at the McMaster University Ancient DNA Centre (Hamilton, ON, Canada). These were:
Abrocoma boliviensis [Museum of Vertebrate Zoology (MVZ) 120238, collected in 1955], Santamartamys
rufodorsalis [American Museum of Natural History (AMNH) 34392, collected in 1912], Diplomys labilis
[Field Museum of Natural History (FMNH) 70101, collected in 1954], Pattonomys occasius (FMNH 84259,
collected in 1958), Pattonomys semivillosus (FMNH 69118, collected in 1953), Toromys rhipidurus (FMNH
87244, collected in 1960), Geocapromys ingrahami (FMNH 5624, collected in 1891), and Plagiodontia aedi-
um (FMNH 63876, collected in 1948). For Plagiodontia, an additional fresh-tissue specimen [University of
Vermont, Zadock Thompson Natural History Collections (ZTNH) 843] was sequenced for vWF and RAG1
and combined with cyt-b and GHR fragments from FMNH 63876 for analysis. We also included additional
DNA sequences from dried tissues of Olallamys albicauda, Toromys grandis, Isothrix barbarabrownae, I.
orinoci, and I. pagurus reported by Upham et al. (2013).
For aDNA specimens, dried tissues adhering to the cranium, mandible, and vertebrae (“crusties”)
were removed, shipped at ambient temperature to the Royal Ontario Museum (Toronto, Canada), and
hand-carried to McMaster University. We used published aDNA protocols (Poinar et al., 2006; Debruyne et
al., 2008) at McMaster, including the use of dedicated clean-room facilities for all pre-PCR work. Approxi-
mately 25 mg of each sample along with one extraction blank were incubated in 0.5 ml of 0.5 M EDTA pH
8.0 (EMD Chemicals) for 24 hours at 25º C using gentle agitation (1000 rpm). Samples were centrifuged at
high speed and the EDTA supernatants were transferred to 5 ml tubes and processed separately. To the
remaining tissue in each sample, we added 0.5 ml of custom digestion buﬀer [20 mMTris (EMD Chemi-
cals) at pH 8.0, 0.5% N-Lauroylsarcosine (Sigma), 250 mg/ml Proteinase K (Thermo Fisher Scientiﬁc), 5
mM CaCl2 (EMD Chemicals), 50 mM DTT (EMD Chemicals), 1% PVP (EMD Chemicals), and 2.5 mM PTB
(Prime Organics)]. After incubating at 55°C for 12-24 hours on a rotary wheel, samples were again spun
down and the supernatant transferred to the same 5 ml tube. This demineralization-digestion process
was repeated three times and the resultant ~3 ml of supernatant formed the raw DNA extract for further
puriﬁcation. Leftover tissue pellets were discarded or saved for future use. Organic puriﬁcation was per-
formed on each supernatant sample, ﬁrst using 1.0 ml of phenol-chloroform-isoamylalcohol (25:24:1, pH
8, Fluka) and then 1.0 ml of chloroform (Fluka), retaining only the aqueous phase in each step. Samples
were concentrated by ultraﬁltration with Amicon Ultra 30K columns (Millipore), washed with three steps
of 450 l of 0.1x TE+Tween [10 mMTris, 0.1 mM EDTA, 0.05% Tween-20 (Sigma), pH 8.0], and the DNA was
ﬁnally eluted in 80 µL of 0.1x TE+Tween.
Polymerase chain reaction (PCR) was carried out on all DNA extractions to amplify target genes. Each
PCR had a reaction volume of 10 l and contained 1.0 l of DNA template, 1.0 l 10x reaction buﬀer, 1.0
l of 8 mM premixed deoxynucleotide triphosphates (dNTPs; 200 M each nucleotide in ﬁnal reaction),
1.0 l of 25 mM MgCl2, 0.5 l of 10 mg/l bovine serum albumin (Applied Biosystems), 4.4 l of double-
distilled H2O (dH2O), 0.1 l of 5 U/lAmpliTaq Gold™ DNA polymerase (Applied Biosystems), and 0.5 l
of each 10 M priming oligonucleotide. PCRs performed on aDNA samples diﬀered from fresh DNA
reactions in the concentration of dNTPs (400 M) and DNA polymerase (0.2 U), the targeting of smaller
gene fragments (200-400 bp), and the inclusion of DNA extraction blanks. All PCR primers, primer pairs,
reaction proﬁles, and DNA sequencing details are given in Upham et al. (2013). Sequences were edited
and assembled using Geneious 6.1.6 software (Biomatters).
Upham and Patterson
Sequence alignment and gene combinability
Of the five genes analyzed, cyt-b was sampled from all but 10 of 199 total taxa (95% complete),
while the sampling of other gene alignments was less complete (12S rRNA, 58%; GHR, 47%; vWF,
42%; and RAG1, 32%; App. 1).Two taxa were represented by chimeras in order to sample all five
genes: Pedetes (union of P. capensis and P. surdaster) and Ctenodactylus (union of C. gundi and C.
vali). All other taxa were single species, although in many cases, they were composites of genes
from multiple voucher specimens. The use of museum voucher specimens as tissue sources per-
mits these sequences to be reassessed, re-evaluated, and possibly even re-identified with future
Sequences from each gene were multiply aligned to establish character homology in relation
to outgroups. For 12S rRNA, we aligned sequences based on the secondary structural model of
Springer and Douzery (1996) after an initial alignment generated in MUSCLE (Edgar, 2004). At sites
where multiple indels made sequence alignment ambiguous, we discarded a total of 198 base
pairs (bp) from the initial alignment of 1035 bp (positions 70-93, 117-125, 219-236, 299-302, 326-
333, 384-408, 426-437, 503-515, 711-719, 782-804, and 934-986). Alignment trials with Gblocks
v0.91b (Castresana, 2000) for 12S rRNA using the same settings as recent studies of rodent phy-
logeny (Montgelard et al., 2008; Fabre et al., 2012a) failed to remove several ambiguous regions, so
these results were discarded. For protein-coding genes, we aligned sequences using ClustalW 2.1
(Larkin et al., 2007) and verified that indels were in sets of three bp. Details of our resulting align-
ments are listed in Table 2.
We explored the potential for mutational saturation in all genes, particularly cyt-b (Griﬃths, 1997; Farias et
al., 2001), using Xia’s test implemented in DAMBE v5.3.52 (Xia et al., 2003; Xia, 2013). Xia’s test compares a stan-
dard index of substitution saturation (Iss) with an index of critical substitution saturation (Iss.c) calculated from
the data via a resampling process to conservatively assess the degree of saturation in a gene. We ran tests for all
genes using all sites, 200 replicates, and specifying the proportion of invariant sites. We also visually examined
cyt-b saturation with plots of pairwise transitions and transitions versus genetic distance on a per codon basis
and over all sites (see App. 3).
Prior to combining gene alignments, we explored the possibility of incongruence between
gene histories (Wiens, 1998). To compare gene-tree topologies most accurately among differen-
tially sampled data sets, we pruned each alignment to the most inclusive set of 50 species that were
sampled for all five genes. Identical 50-species analyses were also performed on mtDNA (cyt-b + 12S
rRNA) and nuclear exon (GHR + vWF + RAG1) data sets to compare phylogenetic patterning among
genome sources with different routes of inheritance. Maximum likelihood (ML) phylogenetic trees
were constructed from each data set using RAxML-HPC2 v7.6.3 (Stamatakis, 2006) on the XSEDE
online computing cluster accessed via the CIPRES Science Gateway (Miller et al., 2010). The best-fit
model of nucleotide evolution for each gene was general time-reversible (GTR) plus among-site
rate variation (G), as found using jModelTest v2.1.4 (Darriba et al., 2012). Models including G and
the proportion of invariant sites (I) were in some cases better fitting, but concerns over the non-
independence of I and G (Sullivan et al., 1999; Mayrose et al., 2005; Stamatakis, 2006) motivated us
to employ the simpler GTR+G model in all cases. Rapid bootstrapping was performed for each gene
alignment using the “–f a” option and 1000 bootstrap replicates, resulting in best-scoring ML trees
annotated with values for bootstrap support (BS). We then compared topologies and nodal support
values across data sets (Fig. 2; Tab. 3). The absence of topological conflict at basal nodes (Tab. 3;
see Results for discussion) allowed us to concatenate the single-gene data sets into a superma-
trix of characters for all 199 taxa, thereby maximizing both taxonomic and genetic diversity in the
phylogeny (approach reviewed by de Queiroz and Gatesy, 2007). The presence of more than 2000
characters in the supermatrix for most taxa was expected to override any statistical biases resulting
from missing data (Wiens, 2006; Lemmon et al., 2009).
Caviomorph evolution through time
Table 2. DNA sequence characteristics for each single-gene alignment. Percent missing data was calculated without inter-
nal alignment gaps. Xia’s test compares indices of substitution saturation for each gene alignment such that Iss > Iss.c indi-
cates saturation, Iss < Iss.c indicates no saturation, and a non-signicant dierence (‘NS’) indicates some degree of saturation.
e 5-gene supermatrix was analyzed using ML in RAxML and Bayesian inference (BI)
in MrBayes version 3.2.1 (Ronquist and Huelsenbeck, 2003); both were run on the XSEDE
computing cluster (Miller et al., 2010). Both ML and BI analyses were partitioned using one
DNA partition per gene and the GTR+G model specied, so that model parameters were esti-
mated independently by partition. RAxML runs were executed using the rapid ML search and
bootstrapping options with 1000 replicates and were repeated several times with random start-
ing trees to verify both topology and clade support values (log-likelihood of -96325.2).
MrBayes runs were started with uniform priors and consisted of four concurrent incremen-
tally heated chains (Metropolis-coupled Markov Chain Monte Carlo, MCMC; Ronquist and
Huelsenbeck, 2003), sampling every 1000 generations over 40 million generations each. Six
independent runs from random starting trees (three sets of two runs each) were compared by
plotting -ln likelihood per generation in Tracer v1.5 (Rambaut and Drummond, 2007), and
comparing marginal densities. After discarding the rst 10% of samples as burn-in, convergent
MCMC searches allowed us to combine a total of 216,000 trees (log-likelihood of -97040.3).
We summarized the combined trees in TreeAnnotator v1.7.5 (Drummond and Rambaut,
2007), resulting in a single maximum-clade-credibility tree with the best a posteriori topology
and nodes annotated with Bayesian posterior probabilities (PP).
ĐǇƚͲbϭϮ^ƌZE GHR ǀt& RAG1
No. taxa aligned 189 115 94 83 63
Total length (bp) 1140 837 884 1269 1064
Pars. inform. (all) 632
351 439 568 265
(codons 1, 2, 3)
Missing data 13.70% 8.80% 8.60% 15.10% 16.60%
Invariant sites 31.70% 37.00% 13.40% 25.50% 47.10%
Base freq: A 30.80% 36.30% 29.20% 21.40% 26.10%
C 26.40% 21.60% 25.30% 28.90% 25.70%
G 12.60% 18.70% 22.50% 30.60% 26.80%
T 30.10% 23.40% 23.00% 19.20% 21.50%
Chi-square 577.5 128.5 118.6 158.1 80.8
df 564 342 279 246 186
P 0.338 ~1.00 ~1.00 ~1.00 ~1.00
Xia’s test: saturated NS unsaturated NS saturated
Iss 0.923 0.65 0.454 0.806 0.896
Iss.c 0.757 0.735 0.739 0.764 0.752
P < 0.05 0.0725 < 0.05 0.1325 < 0.05
Upham and Patterson
We assembled and evaluated the use of 25 fossil calibrations (points A-Y in Fig. 3; App.
2) for estimating clade divergence times in Ctenohystrica. ese calibrations were grouped in
eight dierent sets according to taxonomy, crown vs. stem placement, and previous analyses
(Fig. 4.1). Resulting age estimates were then compared across calibrated nodes (Fig. 4.2), with
non-overlapping 95% credibility intervals indicating that estimates were signicantly dierent
between calibration sets. e placement of calibrations relied on recent cladistic studies of living
and fossil taxa (e.g., Antoine et al., 2012; Pérez and Pol, 2012; Verzi et al., 2013) to allocate
fossils among crown or stem groups depending on shared-derived morphological characters.
Appendix 2 lists and justies the minimum and maximum ages for each calibration point with
reference to fossil taxa, locality and stratigraphic levels of collection, geologic age estimates,
and analyses that identify the phylogenetic position of fossils, following recommendations by
Parham et al. (2012). Minimum ages were set as lognormal priors, assuming that the oldest
condently assigned fossil in a group constrains that group’s origination age. Soft maximum
ages were set as the rear 5% of the lognormal distribution using dates from the youngest fossil
assemblage that lacked fossils allied to the calibrated group (see App. 2).
We estimated clade divergence times using the Bayesian relaxed-clock model implemented in
BEAST 1.7.5 (Drummond et al., 2012), and a starting tree obtained from penalized likelihood (PL) analy-
ses in r8s v1.8 (Sanderson, 2003).We used the PL starting tree to meet the age constraints of all fossil priors
and prevent BEAST runs from crashing; subsequent analyses using the MCMC framework in BEAST
allowed for information in the original molecular data to be incorporated into divergence-time estimates,
and for the temporal inuence of dierent fossil priors to be tested. We started the PL analyses using the
5-gene MrBayes tree pruned to 198 taxa (removing Pedetes) to achieve the fully bifurcating tree required by
BEAST. We set PL constraints using the age minima and 95% maxima of all 25 calibrations (App. 2), with
the exception of the Ctenohystrica root, which we xed to the 50 Ma median age of its prior (r8s requires
one xed point). We arbitrarily changed several Ctenomys branch lengths that were near zero to the value
“0.0011” to avoid the need to collapse branches (Sanderson, 2003). Cross-validation in r8s determined an
optimal smoothing parameter of 2.5, and the resulting tree was used to start all BEAST runs.
We focused BEAST runs on estimating divergence times rather than topology by constraining
the monophyly of all clades recovered from the MrBayes analysis (except those within Ctenomys).
We ran BEAST analyses under the GTR+G model with four gamma categories, unlinking site
models across the ve gene partitions and estimating base frequencies. Relaxed clock models were
unlinked except for the mtDNA genes (linked on the same strand) and rates were uncorrelated
so that each branch was estimated from independent draws of a lognormal distribution. e uni-
versal clock mean priors were set to uniform with a large upper bound. Tree models were linked
and the tree prior was set to Yule for the trial analyses with calibration sets 1-8. e nal analysis
used the birth-death tree prior with incomplete species sampling (Stadler, 2009), given 0.69 as the
fraction of extant ctenohystricans sampled. MCMC chain lengths were set to 10 million genera-
tions with parameters sampled every 10,000 generations. For all analyses, eight independent runs
were performed on the XSEDE computing cluster and combined after 10% burn-in, resulting in
Caviomorph evolution through time
7,200 trees (72 million generations) that in each case converged upon stable posterior distribu-
tions (determined using Tracer). Trees were summarized with the “maximum clade credibility”
option using TreeAnnotator. Timetrees containing mean divergence times and 95% credibility
intervals for each node were compared in R using the ape and phyloch packages (Paradis et al.,
2004). Below we report mean divergence times, but invite inspection of the associated error bars,
which vary in width given the proximity and shape of age priors (Tab. 4 and Fig. 5).
DNA sequence analyses
Table 2 contains DNA sequence characteristics for each of the ve single-gene alignments.
Although no parsimony analyses were conducted here, the number of parsimony-informative
sites (PIS) per locus is an instructive metric of raw sequence variation. e cyt-b alignment
contains more PIS than any other sampled gene, owing to its extensive taxonomic sampling and
greater variability. Correcting for sites and taxa, vWF contains the most PIS (0.0054), followed
by GHR, RAG1, 12S rRNA, and cyt-b (0.0053, 0.0040, 0.0036, and 0.0030, respectively).
Both mtDNA genes contain fewer G nucleotides than do the nuclear exons, but no signicant
dierences are found in nucleotide composition within each gene (P> 0.05; Tab. 2).
To verify aDNA results, we repetitively amplied each gene fragment and assembled multiple
overlapping fragments for each gene (Pääbo et al., 2004). Because instances of polymorphism
among nuclear DNA fragments could reect either DNA damage or diploid heterozygosity, we
coded all polymorphic sites with the corresponding IUPAC ambiguity codes.
Results from Xia’s test for substitution saturation show that four of the ve sampled genes
display some degree of saturation (Tab. 2). e cyt-b and RAG1 alignments each display sig-
nicant saturation and only the GHR alignment is found to be signicantly unsaturated. Plots
of the pairwise number of transitions and transversions in cyt-b versus percent sequence diver-
gence conrm substitution saturation across the breadth of the tree (App. 3). Given that cyt-b’s
rapid evolution makes it a useful species-delimitation marker (e.g., Baker and Bradley, 2006),
saturation was actually expected at this taxonomic scale. Cyt-b nonetheless provides important
resolution at terminal nodes (Tab. 3). Finding substitution saturation in 12S rRNA, vWF, and
RAG1 was, however, unexpected, but follows from the Eocene-Recent range of divergences in
each data set, especially given the conservative design of Xia’s test (Xia and Lemey, 2009). We
chose to proceed with caution, testing whether these varying degrees of saturation actually result
in conicting phylogenetic signal by comparing the statistical support for equivalent nodes in
the phylogenies of each data partition.
Single-gene, mtDNA, and exon data sets: completely sampled trees
Results from comparisons of the 50-taxon trees, each completely sampled for all ve
genes, are displayed in Table 3 with node numbers from Figure 2. Across 48 total nodes, only
three instances of signicant conict in topology are found among the single-gene data sets,
each of which occurs at shallow nodes (Tab. 3). Because these conicts exist among closely re-
lated taxa and involve support for alternative sister-relationships among species (node 42) or genera
Upham and Patterson
Figure 2. Phylogeny based on the concatenated analysis of 5 genes (cyt-b, 12S rRNA, GHR, vWF, and RAG1) that were com-
pletely sampled for 50 species representing all major groups of Ctenohystrica (Bayesian inference topology shown). Node
numbers in this gure dene groupings of taxa that facilitate comparisons of topology across single gene and concatenated
gene data sets; support values for these permutations are displayed in Table 3. Thick branches indicate nodes with Bayesian
inference posterior probability (PP) ≥ 0.95 and maximum likelihood bootstrap support (BS) ≥ 85%.
Caviomorph evolution through time
Table 3. Nodal support values for single genes and combined gene data sets constructed using only the 50 taxa that were
sampled for all ve genes. Node numbers are listed with reference to Figure 2. All values are maximum-likelihood bootstrap
percentages (ML BP) except the column headed BI PP (Bayesian inference posterior probability). Boldface values indicate ML
BP ≥ 85 or BI PP ≥ 0.95. Dierences among data sets are marked as “-” if ML BP < 85, and with one or more asterisks (*) if they
are statistically supported by ML BP ≥ 85 (see Table footnotes and Results section for further details).
ϱͲŐĞŶĞ ϮͲŐĞŶĞ ϯͲŐĞŶĞ
EŽĚĞ /WW D>W ŵƚE ĞǆŽŶƐ ĐǇƚͲbϭϮ^ƌZE GHR ǀt& RAG1
1ϭ͘ϬϬ ---- - ---
2ϭ͘ϬϬ ϭϬϬ 83 ϭϬϬ 46 47 ϭϬϬ 88 98
3Ϭ͘ϵϵ 77 47 58 - 33 63 - 41
4ϭ͘ϬϬ ϭϬϬ 68 99 -52ϭϬϬ --
5ϭ͘ϬϬ ϭϬϬ 98 ϭϬϬ 75 93 ϭϬϬ ϭϬϬ 49
6ϭ͘ϬϬ 98 52 97 -7398 -48
7ϭ͘ϬϬ ϭϬϬ 77 ϭϬϬ 54 - 94 98 -
8ϭ͘ϬϬ ϵϬ 31 88 ---6634
9ϭ͘ϬϬ ϭϬϬ 95 ϭϬϬ -ϵϬ ϭϬϬ ϭϬϬ ϭϬϬ
10 ϭ͘ϬϬ 83 66 - 58 26 88 --
11 ϭ͘ϬϬ ϭϬϬ 84 99 77 48 91 -95
12 ϭ͘ϬϬ ϭϬϬ 98 ϭϬϬ 93 80 ϭϬϬ 91 97
13 ϭ͘ϬϬ 81 91 -7581 *85 -
14 ϭ͘ϬϬ ϭϬϬ 94 ϭϬϬ 84 23 ϭϬϬ 98 ϭϬϬ
15 0.50 - - - - - - 29 -
16 ϭ͘ϬϬ 99 22 97 ---81-
17 ϭ͘ϬϬ 97 46 79 - 45 57 - -
18 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ 99 98
19 ϭ͘ϬϬ 91 97 -99 -63--
20 ϭ͘ϬϬ ϭϬϬ ϭϬϬ 98 92 99 97 89 75
21 0.52 46 29 - - - - - -
22 0.78 48 45 42 37 23 - - -
23 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ
24 ϭ͘ϬϬ ϭϬϬ 82 ϭϬϬ 55 73 92 85 -
25 ϭ͘ϬϬ 83 94 -86 - - 35 44
26 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ 62 96 96 -
27 Ϭ͘ϵϱ 72 68 - 69 71 61 - -
28 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ 97 99 99 77
29 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ 86
30 ϭ͘ϬϬ ϭϬϬ 29 99 - - 82 65 28
31 ϭ͘ϬϬ 97 66 55 53 - 47 - -
32 ϭ͘ϬϬ ϭϬϬ 42 ϭϬϬ --ϭϬϬ ϭϬϬ 91
33 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ 96 96 94 ϭϬϬ 98
34 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ 98 94 99 99 ϭϬϬ
35 0.88 60 30 - 31 - - - -
36 0.52 - - 36 - - - 34 45
37 Ϭ͘ϵϲ 77 - 68 - - - - 40
38 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ 88 67
39 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ ϭϬϬ 92 ϭϬϬ ϭϬϬ 93
40 ϭ͘ϬϬ ϭϬϬ 97 ϭϬϬ 92 ϵϬ ϭϬϬ 99 85
41 ϭ͘ϬϬ 81 65 72 52 47 91 --
42 ϭ͘ϬϬ ϭϬϬ ϭϬϬ 80 ϭϬϬ 63 97 ** -
43 ϭ͘ϬϬ ϭϬϬ ϭϬϬ ϭϬϬ 77 99 98 98 76
44 ϭ͘ϬϬ 98 64 92 -74-95 -
45 ϭ͘ϬϬ ϭϬϬ 67 ϭϬϬ 45 - 97 99 75
46 ϭ͘ϬϬ 80 76 36 78 - - - 79
47 ϭ͘ϬϬ ϭϬϬ 95 ϭϬϬ 93 -ϭϬϬ 97 99
48 0.93 - 72 - 97 - - 65 ***
* GHR shows 86% support for Octodontomys – Salinoctomys relationship
** vWF shows 88% support for Mesocapromys angelcabrerai – Mysateles prehensilis relationship
*** RAG1 shows 88% support for Hydrochoerus – Galea relationship
Upham and Patterson
5-gene data sets: completely sampled versus supermatrix trees
We compared the topology from the 5-gene completely sampled tree (CST; Fig. 2) with
the positions of the same taxa in an analysis based on a supermatrix of all 199 taxa collapsed
to genera (Fig. 3). Both analyses strongly support Hystricognathi (Old World porcupines,
phiomorphs, and caviomorphs; node 2), but only the supermatrix strongly supports Hystricidae
as sister to Phiomorpha + Caviomorpha (node 3). Both Phiomorpha and Caviomorpha are
strongly recovered as monophyletic in both analyses (nodes 4 and 7). Within phiomorphs,
strong evidence supports Bathyergidae (node 6) and Petromuridae + ryonomyidae (node
5) in both sets of analyses. Within caviomorphs, both sets of analyses securely recover the
four recognized superfamilies as monophyletic (nodes 45, 43, and 9) and forming sister pairs:
Erethizontoidea + Cavioidea and Chinchilloidea + Octodontoidea (nodes 44 and 8).
At the family level, there is strong or moderate support for the monophyly of the
caviomorph families Erethizontidae, Cuniculidae, Dasyproctidae, Caviidae, Dinomyidae,
Chinchillidae, Abrocomidae, Octodontidae, Ctenomyidae, and Capromyidae. e CST (Fig.
2) loosely supports Cuniculidae + Dasyproctidae (node 46), whereas this group is moderately
supported in the supermatrix analysis. Dinomyidae + Chinchillidae (node 43) are strongly
supported as a clade in both analyses. Abrocomidae appears as the sister to all other octodontoid
families (node 10), with strong support for its sisters, Ctenomyidae + Octodontidae (node
11) and Capromyidae + Echimyidae (node 14). However, whereas the CST analysis recovers a
monophyletic Echimyidae (node 15), the supermatrix analysis nds Echimyidae paraphyletic
with respect to Capromyidae (node 39); a group of eastern Brazilian taxa (Carterodon, Clyomys,
Euryzygomatomys, and Trinomys) are loosely recovered as sister to the capromyids (Fig. 3).
At the genus level, there is also strong agreement between analyses in the recovered
relationships. ese include Salinoctomys (Octodontomys, Octodon); Geocapromys (Capromys
(Mysateles, Mesocapromys); Trinomys (Clyomys, Euryzygomatomys); Myocastor (Hoplomys,
Proechimys); and Makalata (Echimys, Phyllomys). However, minor dierences are apparent in
the topologies for a handful of genera. e supermatrix oers support for Hydrochoerus (Cavia,
Galea), whereas the CST loosely recovers Galea as sister to Cavia + Hydrochoerus (nodes 47 and
48, Fig. 2). Mesomys and Isothrix form a moderately supported clade in the supermatrix, while
in the CST they loosely appear as successive sisters to the remaining arboreal echimyids (nodes
17 and 21, Fig. 2). Callistomys and Myocastor appear as either successive sisters to Hoplomys +
Proechimys (supermatrix) or as a poorly supported clade that is sister to them (CST).
(nodes 13, 48), they are likely the result of incomplete lineage sorting. In the absence of topo-
logical conict at deeper nodes between single-gene trees, we concatenated data sets into 2-gene
(mtDNA), 3-gene (exon), and 5-gene (combined) data sets. As expected, the greater degree of
saturation in the mtDNA data set results in poor support at most basal nodes in the phylogeny
(nodes 1-25 in Fig. 2) and higher nodal support among terminal branches, but there is no
signicant conict between genes inherited maternally (mtDNA) and bi-parentally (nuclear
exons; Tab. 3). e exon data set shows high resolution at basal nodes in the tree as well as at
terminal branches, and agrees with each of the other data sets. e node-by-node concordance
of all phylogenies from the 50-taxon data sets justies the concatenation of data frames for all
199 taxa sampled, resulting in a 5-gene supermatrix.
Caviomorph evolution through time
Figure 3. Genus-level phylogeny of Ctenohystrica from the 5-gene supermatrix (cyt–b + 12S rRNA + GHR + vWF + RAG1) focus-
ing on the South American rodent clade Caviomorpha and including sister clades from Africa and Asia (Phiomorpha, Hystricidae,
Diatomyidae, and Ctenodactylidae). The gene sampling matrix on the right is coded to show the sampling of genes for one or
more species in a genus and the origin of the DNA sequences whether produced for the present study (black boxes), deposited
on GenBank from previous studies (grey boxes), or currently unsampled (white boxes). Sequences from Upham et al. (2013) are
included as black boxes. A total of 68 genera are represented in this phylogeny (Bayesian topology shown). Nodes are annotated
with support values as indicated from Bayesian inference posterior probability (PP) and maximum likelihood bootstrap support
(BS) analyses. Asterisks (*) denote instances where nodes were recovered in the Bayesian but not in the maximum likelihood analy-
ses. Letters in boxes correspond to fossil calibration points, as detailed in Appendix 2. See text for details on lineages I, II, and III
Upham and Patterson
Only the supermatrix analysis (Fig. 3) had the taxonomic scope to resolve a number of addi-
tional generic relationships. Within Hystricidae, there was strong support for Trichys (Atherurus,
Hystrix). Within Bathyergidae, there was strong support for Heterocephalus as sister to all other
mole-rats, followed by Heliophobius, and the remaining mole-rats form a monophyletic group-
ing of Georychus + Bathyergus and Cryptomys + Fukomys. Within Caviidae, Dolichotis is sister
to Hydrochoerus + Kerodon, and this group is sister to Galea (Microcavia, Cavia). In Chinchilli-
dae, Lagostomus is sister to Chinchilla + Lagidium. Abrocoma and Cuscomys are both placed in
Abrocomidae. Octodontidae is recovered as two well-supported clades: Octomys (Pipanacocto-
mys (Salinoctomys, Tympanoctomys) and Octodontomys (Octodon (Spalacopus, Aconaemys). Within
Echimyidae, there was strong support for –Lonchothrix + Mesomys; ((Santamartamys, Diplomys)
(Olallamys, Kannabateomys, Dactylomys)), although the latter trichotomy could not be con-
dently resolved; and a lineage of arboreal rats comprised of two subclades, Pattonomys + Toromys
and Makalata (Echimys, Phyllomys).
Timetree calibration trials
e eight sets of fossil calibrations (Fig. 4.1) form three groups that are statistically
equivalent in terms of overlapping 95% credibility intervals on all calibrated nodes. ese are
group A, including sets 1, 4, and 6; group B, including sets 2, 3, and 8; and group C, including
sets 5 and 7 (Fig. 4.1). Considering pairwise dierences in sets across the 25 calibrated nodes,
sets in group A nd signicantly dierent ages on an average of 8 nodes and 7 nodes from sets
in groups B and C, respectively (ranges 4-13). e group A sets dier in number of calibrations
(25, 20, or 7), but have in common the 7 stem fossils in set 6 (Fig. 4.1). e calibrations O and
T are unique to the group A sets, and therefore appear to be responsible for the uniformly older
age estimates in these sets versus those in groups B and C (Fig. 4.2). Excluding calibrations O
and T (set 7) nds results that are equivalent to using 18 crown calibrations (set 5). ese group
C sets dier from those in group B on the age estimate for 1 calibrated node: U, the calibration
for crown Capromyidae (~15 Ma vs. ~10 Ma; Fig. 4.2).e older age of node U when this
node is calibrated highlights uncertainty regarding this fossil placement (MacPhee et al., 2003),
motivating us to exclude this calibration from further analyses (see also the “Capromyidae”
section in the Discussion). We thus selected set 8 –excluding calibrations O, T, and U– for the
nal analyses. Set 8 provides estimates that are equivalent to sets with fewer calibrations (the
group B sets 2 and 3), but because it retains information from 22 calibrations it does so with the
most precision of any set evaluated (mean 95% credibility interval of 4.3 Ma, versus 5.9, 5.8,
4.8, 7.0, 5.5, 8.3, and 4.4 Ma for sets 1-7).
e divergence time analysis using calibration set 8 (log-likelihood of -93143.8) presents
a calibrated chronology of group appearance within the Ctenohystrica (Fig. 5). Hystricognathi
had appeared by roughly 50 Ma, and by 44.9 Ma, the Old World porcupines had diverged
from them. e split between phiomorphs and caviomorphs is dated at 42.0 Ma, with the
most recent common ancestor (MRCA) of phiomorphs dated to 36.3 Ma and the MRCA of
caviomorphs to 35.7 Ma. Subsequent splits in these lineages took place in the early Oligocene;
Caviomorph evolution through time
Figure 4. Comparison of eight sets of fossil calibrations that dier in their 1. inclusion of 25 calibrated nodes (AY), and 2.
resulting inuence on mean age estimates (red) and 95% credibility intervals (blue). Only the 15 calibrated nodes with age
estimates that diered signicantly among calibration sets are displayed; the other ten calibrated nodes had overlapping
95% credibility intervals across all sets. Asterisks (*) on plots for calibrations O, T, and U indicate that their inclusion in sets
1, 4, and 6 yielded results that were inconsistent with other sets. Exclusion of these three calibrations yielded the most
consistent age estimates (set 8, in bold); results from this set of 22 calibrations are depicted in Figure 5 and discussed in
the text. Nodes A–Y are labeled in Figure 3. Groups A, B, C indicate statistically equivalent sets (see Results). See Appendix
2 for calibration details and justications.
the MRCAs of Diatomyidae + Ctenodactylidae (32.2 Ma), Bathyergidae (31.2 Ma), and the
near-simultaneous divergences of Erethizontoidea + Cavioidea (32.4 Ma) and Chinchilloidea +
Octodontoidea (32.8 Ma) all date to this interval. e crown radiations of Cavioidea (25.2 Ma)
and Octodontoidea (25.6 Ma) were both initiated by the end of the Oligocene, followed closely
by their subdivision into the cavioid families Cuniculidae, Dasyproctidae, and Caviidae (23.4
Ma), and the octodontoid family Abrocomidae and dyads of Octodontidae-Ctenomyidae and
Echimyidae-Capromyidae (23.6 Ma).e two other caviomorph superfamilies have MRCAs
during the early-to-mid Miocene: 17.5 Ma (Erethizontoidea) and 19.7 Ma (Chinchilloidea).
Other family and genus divergence timings are detailed in Table 4 along with a comparison of
analogous results from other studies. Overall, our targeted eort to reconstruct the Ctenohystrica
Upham and Patterson
ůĂĚĞ dŚŝƐƐƚƵĚǇ hƉŚĂŵĂŶĚ DĞƌĞĚŝƚŚet al͘ZŽǁĞet a l͘ ^ĂůůĂŵet al͘ ,ƵĐŚŽŶet al͘
WĂƩĞƌƐŽŶ;ϮϬϭϮͿ ;ϮϬϭϭͿ ;ϮϬϭϬͿ ;ϮϬϬϵͿ ;ϮϬϬϳͿ
Ctenohystrica 50.0 (46.2-55.0) 50.8 (41.0–69.7) 61.1 (56.1–68.3) 92 (68–121) 55.6 (52.7–58.3) 61.3 (53.8–63.3)
Diato-Cteno 32.2 (28.5–37.6) b43.1 (38.2–49.0) b7.9 (5.2–12.1) 44.3 (37.6–51.1)
Ctenodactylidae 13.4 (7.3–20.5) aaaa
Hystricognathi 44.9 (42.5–47.4) 43.7 (37.2–51.0) 49 (41.9–56.7) 59 (44–74) 39.0 (36.1–41.9) 45.4 (39.7–50.5)
Hystricidae 12.7 (11.0–15.8) aaa
15.7 (11.7–21.1) a
Phiomorpha / Caviomorpha 42.0 (41.1–43.3) 40.4 (36.1–45.6) 47.2 (39.8–55.1) c36.1 (33.4–39.0) 42.7 (37.3–47.2)
Phiomorpha 36.3 (32.6–39.4) 36.3 (30.9–42.5) 42.0 (35.7–49.7) 55 (41–70) 32.1 (29.2–35.1) 38.0 (32.7–42.9)
Petro-Thryo 23.0 (17.6–28.1) 23.5 (17.5–29.5) 26.2 (18.3–33.3) 34 (22–49) 18.0 (14.8–21.3) 24.5 (19.8–42.9)
Bathyergidae 31.2 (26.7–35.1) 31.5 (25.3–37.4) a~46 26.3 (22.7–29.7) 32.8 (27.6–38.0)
Bathyergidae minus Heter 17.9 (14.3–21.8) 18.6 (13.6–23.4) b~15 aa
Caviomorpha 35.7 (33.8–37.6) 34.1 (36.1–45.6) 42 (34.0–49.0) 45 (35–54) 30.1 (27.9–33.4) 34.3 (29.9–36.9)
Erethizontoidea /Cavioidea 32.4 (31.4–33.8) 30.6 (27.4–34.7) 40.9 (33.3–47.9) c28.6 (25.9–31.4) 31.7 (27.4–35.1)
Erethizontoidea 17.5 (11.3–24.0) 7.5 (5.0–10.4) a15 (8–23) aa
Cavioidea 25.2 (24.3–26.7) 30.6 (27.4–34.7) 32.2 (25.0–38.0) 37 (28–47) 23.1 (20.6–26.0) 24.1 (20.1–28.0)
Cunic-Dasyp 23.4 (21.0–25.7) cccbb
ĂƐǇƉƌŽĐƟĚĂĞ 8.1 (6.0–10.7) 10.2 (6.6–14.7) a9 (5–16) bb
Caviidae 17.6 (14.9–20.6) 19.2 (16.0–22.7) 19.9 (14.8–25.7) ~25 14.6 (12.4–17.2) a
Chinchilloidea / Octodontoidea 32.8 (31.4–34.5) 32.7 (30.3–36.4) 38.9 (31.3–46.2) 43 (33–52) 29.0 (26.2–31.7) 33.3 (28.9–36.3)
Chinchilloidea 19.7 (14.8–24.7) 19.0 (13.7–25.1) 28.2 (24.3–34.7) 29 (20–38) 19.9 (16.5–23.2) 21.4 (17.9–25.6)
Chinchillidae 12.3 (9.3–15.9) 14.3 (8.1–21.2) a~24 aa
Octodontoidea 25.6 (23.1–27.9) 26.8 (24.8–28.9) 28.2 (21.0–33.7) ~26 19.6 (17.1–22.2) b
Abrocomidae 8.4 (7.2–15.0) 0.3 (0–1.8) aaab
Octod-Cteno / Capro-Echim 23.6 (21.4–25.8) 25.3 (24.6–26.7) 26.7 (19.6–31.8) ~25 18.1 (15.7–20.6) 18.6 (15.3–21.8)
Octod-Cteno 18.9 (15.7–22.1) 19.1 (14.3–23.5) 22.9 (17.1–27.2) 22 (17–30) bb
KĐƚŽĚŽŶƟĚĂĞ 8.8 (7.3–10.4) 9.0 (6.7–11.6) a8 (5–12) aa
Ctenomyidae 6.0 (4.6–7.6) 4.3 (2.2–7.4) aabb
Capro-Echim 18.2 (17.1–19.3) 18.8 (17.7–20.6) 20.0 (13.7–24.0) 15 (11–21) 12.3 (9.9–14.7) 12.4 (9.7–15.3)
Capromyidae 9.8 (7.4–12.3) aaaaa
Echim: clade 1 16.7 (15.1–18.1) 16.4 (13.8 18.8) bbbb
Echim: clade 1-clade 2 17.1 (16.4–18.1) 16.6 (14.9–18.4) b~13 bb
Echim: clade 2 15.5 (14.2–16.8) 14.9 (13.0–16.9) b~11 bb
Echim: clade 3 16.2 (15.8–16.9) 15.6 (13.9–17.6) b~12 bb
Table 4. Comparison of divergence times among studies. All times are in millions of years and refer to the estimated ages (and condence intervals) of the specied crown groups.
Results from this study are from the timetree in Figure 4.
Abbreviations. Cteno, Ctenodactylidae; Cunic, Cuniculidae; Dasyp, Dasyproctidae; Diato, Diatomyidae; Heter, Heterocephalus; Petro, Petromuridae; Thryo, Thryonomyidae. Upham and Pat-
terson (2012) analyzed 12S rRNA, GHR, vWF, and RAG1 for 51 ctenohystrican genera; Meredith et al. (2011) analyzed 26 gene fragments across mammalian families, including 19 genera in
Ctenohystrica (“global mean” and “DNA all” analyses); Rowe et al. (2010) analyzed GHR and TTR for 39 ctenohystrican genera (U=55 Ma and “All/gap” analyses); Sallam et al. (2009) analyzed 115
morphological characters from Marivaux (2004) and ADRA2B, IRBP, vWF, GHR, RAG2, and CB1 from Blanga-Kanﬁ et al. (2009) for 18 ctenohystrican genera; and Huchon et al. (2007) analyzed
ADRA2B, IRBP, vWF, GHR, cyt-b, and 12S rRNA for 16 ctenohystrican genera.
aOnly one taxon sampled (no crown age); bOne or more taxa not sampled in analysis; and cAnalysis did not recover sister relationship.
Caviomorph evolution through time
timetree resulted in a greater breadth of sampled lineages and narrower nodal error bars than
other recent timetrees (Tab. 4). Age estimates from the present study are comparable to
our earlier study using ve fossil calibrations (Upham and Patterson, 2012), but improved
taxon sampling enables crown ages for several families to be estimated (e.g., Abrocomidae,
Capromyidae, Hystricidae). Prior estimates for the origin of crown Ctenohystrica varied from
~50 Ma (Sallam et al., 2009; Upham and Patterson, 2012) to ~61 Ma (Huchon et al., 2007;
Meredith et al., 2011) all the way to the mid-Cretaceous ~92 Ma (Rowe et al., 2010).
Appraisals of rodent evolution have taken enormous steps forward in recent years. e
analysis of molecular datasets has oered a means to sidestep the morphological conservatism
and parallelism (e.g., Wood, 1935) that have plagued rodent systematics for more than a century
(see also Castoe et al., 2009). Comprehensively sampled molecular phylogenies across Rodentia
(e.g., Honeycutt, 2009; Fabre et al., 2012a) and the present eort for hystricognath rodents now
permit a fuller appraisal of rodent evolutionary and biogeographic diversication through time.
The Ctenohystrica and higher-level relationships
Our tree unambiguously recovers a monophyletic Hystricognathi, with Hystricidae sister
to Caviomorpha + Phiomorpha sensu stricto (i.e., Bathyergidae, Petromuridae, and ryono-
myidae). e same set of relationships was uncovered by the analyses of Adkins et al. (2001),
Huchon et al. (2002), Huchon et al. (2007), Honeycutt (2009), Fabre et al. (2012a), and Voloch
et al. (2013). Previous analyses had either regarded this as a trichotomy (Huchon and Douzery,
2001) or recovered Hystricidae as sister to Caviomorpha with Phiomorpha sister to this group
(also called “Bathy-Phiomorpha”; Vilela et al., 2009; Rowe et al., 2010).
Our trees robustly recover the sister-group relationship of Petromuridae with ryono-
myidae (ryonomyoidea), and this group as sister to Bathyergidae. Nedbal et al. (1994) and
Huchon and Douzery (2001) documented this same topology, as have others more recently
(Sallam et al., 2009; Rowe et al., 2010; Meredith et al., 2011).
One remarkable feature of the timetree is the very early divergence of the lineage leading
to Heterocephalus from other mole-rats (Fig. 5), estimated at 31.2 Ma. Contemporaneously, on
another continent, the early Oligocene witnessed the emergence of erethizontoids and cavioids
(32.4 Ma) and of chinchilloids and octodontoids (32.8 Ma). e divergent gene sequences
and early divergence time of Heterocephalus were noted in prior molecular studies (Allard
and Honeycutt, 1992; Janecek et al., 1992; Walton et al., 2000). Currently, Heterocephalus
is considered to be the lone member of the bathyergid subfamily Heterocephalinae (Woods
and Kilpatrick, 2005). However, its morphological distinctiveness and early divergence from
other mole-rats suggest that it should be elevated to family rank within the Bathyergoidea, i.e.,
Heterocephalidae Landry, 1957 (Patterson and Upham, 2014).
Upham and Patterson
Figure 5. Molecular timetree of ctenohystrican rodent evolutionary history calibrated with 22 fossil priors in BEAST (see App.
2). Numbers above nodes are the mean divergence ages of clades, and node error bars correspond to 95% credibility inter-
vals. Black dots on nodes correspond to MrBayes posterior probability (PP) values ≥ 0.95. The timing of geological epochs is
from Gradstein et al. (2004). See text for details on lineages I, II, and III within Echimyidae.
Caviomorph evolution through time
The Phiomorpha-Caviomorpha split
Our phylogeny condently recovers a sister-group relationship between the phiomorph
rodents endemic to Africa (ryonomyidae, Petromuridae, and Bathyergidae) and the Ameri-
can caviomorphs. We estimate their divergence time at 42.0 Ma (41.1-43.3), in the Middle Eo-
cene (Fig. 5). is date is robust to recalculation excluding the fossil assemblage at Contamana,
Peru (calibrations C and E in Fig. 3), yielding ages that are slightly younger, but not signicantly
so, for the phiomorph-caviomorph split (39.9 Ma [36.8-43.2]) and the Hystricognathi crown
(43.2 Ma [39.3-47.1]; modied from calibration set 8). at Peruvian locality contains the
oldest known hystricognath fossils in strata constrained to 41.6-40.94 Ma by a combination of
radiometric and biochronological methods (see App. 2; Antoine et al., 2012).
Our Middle Eocene estimate for the common ancestor of African and American hystricog-
naths is generally on par with recent molecular studies using a variety of markers and taxon-
sampling strategies (Tab. 4). Using immunological distances, Sarich (1985) estimated their
divergence at 45-48 Ma, in agreement with the Eocene divergences hypothesized by Woods
(1982) and Wood (1985). An older estimate of ~85 Ma, consistent with vicariance via Afri-
can and South American plate tectonics, was based on the mitochondrial divergence of cane
rats from guinea pigs (Mouchaty et al., 2001).e majority of DNA-based estimates place the
Phiomorpha-Caviomorpha split in the Eocene, ruling out plate-tectonic explanations in favor
of trans-oceanic dispersal to South America via one of three routes: North America, Africa, or an
Australia-Antarctica landbridge (Huchon and Douzery, 2001). Molecular clock-based estimates
vary across the Eocene, from the latest to the earliest stages (~34-56 Ma; Nedbal et al., 1994;
Huchon and Douzery, 2001; Opazo, 2005; Poux et al., 2006; Huchon et al., 2007; Vilela et al.,
2009; Voloch et al., 2013). Several studies used the then-earliest known caviomorph fossil from
the Tinguirirican of Chile (~31 Ma; Wyss et al., 1993) to calibrate the phiomorph-caviomorph
split, yielding estimates of 43-54 Ma (Huchon and Douzery, 2001), 34.0-39.2 Ma (Opazo,
2005), and 38.9-48.5 Ma (Poux et al., 2006) depending on the calibration methods and genes
analyzed. Rowe et al. (2010) instead used this fossil strictly as “dasyproctid indet.” (Wyss et al.,
1993) to calibrate crown Cavioidea, yielding a considerably older date of 42-69 Ma for the stem
divergence of caviomorphs (the uncertainty of this estimate derives from the poorly supported
Caviomorpha-Hystricidae relationship recovered in that analysis). Recently, this Tinguirirican
caviomorph was identied as Andemys termasi, Pan-Dasyproctidae (Bertrand et al., 2012), and
its age was further constrained to 31.6-33.6 Ma (Dunn et al., 2013). e subsequent discovery
of ~41 Ma caviomorph fossils in Peru (Antoine et al., 2012) supports our use of Andemys as an
age constraint inside Caviomorpha, treating it conservatively as a stem member of Cavioidea
(App. 2; Pérez and Pol, 2012).
e most thorough previous eort to calibrate the divergence of Old and New World hystri-
cognaths was the Mammalia-wide study of Meredith et al. (2011).ey used seven fossil calibra-
tions internal to Ctenohystrica (of 82 total for mammals), along with genetic sampling from all
17 extant families, to estimate an age of 39.8-55.1 Ma for the Phiomorpha-Caviomorpha split.
A total of ve calibrations, all placed internal to Ctenohystrica, were used in Upham and Patter-
son (2012) and yielded an estimated split of 36.1-45.6 Ma. Other recent eorts have included
more calibrations external to Hystricognathi and Rodentia, estimating the phiomorph-cavio-
morph divergence as 33.4-39.0 Ma (Sallam et al., 2009) and ~39-47 Ma (Voloch et al., 2013).
Upham and Patterson
Given the range of sampling in taxa and genes and the dierent methods employed, it is remark-
able that nearly all of these molecular clock studies are in close agreement. Without exception,
each of these works and the present one nd evidence in support of an Old World-New World
divergence prior to the Eocene-Oligocene boundary.
Our estimate for the Phiomorpha-Caviomorpha split, 42.0 Ma, approximately coincides
with the Mid-Eocene Climatic Optimum (MECO –see Fig. 5; Zachos et al., 2008), which
occurred at ~41.5 Ma (Bohaty and Zachos, 2003). Given that the MECO event occurred
shortly before the oldest known South American caviomorph fossils ~41 Ma and shortly after
their inferred split from African phiomorphs, these biotic and abiotic events are coincidentally
linked (Fig. 6). However, evidence that the MECO event initiated this biotic dispersal across the
Atlantic Ocean is purely circumstantial, and unlikely to be corroborated because waif dispersals
are by denition rare and idiosyncratic (e.g., via rafts of oating debris; Lavocat, 1969; Martin,
1994a; Houle, 1998). Nevertheless, the scenario of trans-Atlantic dispersal is not altogether
unrealistic. Houle (1998) used a model of paleocurrents and paleowinds for the Eocene Atlantic
Ocean to estimate that oating debris would have crossed this oceanic barrier in ~11 days.
is nding is consistent with studies that small-to-medium-sized mammals can survive 10-15
days without water, and suggests that caviomorph ancestors may have originated in temperate
regions of Africa with discrete wet-dry seasonality (Houle, 1998; Houle, 1999). e MECO
event is known from deep-sea isotope records in the Southern Atlantic and Indian Oceans –the
same region where trans-Atlantic dispersal is hypothesized– and consisted of 4 °C warming in
less than 1 Ma (Bohaty and Zachos, 2003). If the intensity of coastal storms in Africa increased
from ocean warming, then caviomorph ancestors might have traveled a MECO “conveyer” of
sorts, riding coastal debris across the Atlantic to South America.
South American primates also share an African or Asian ancestor (Kay et al., 1998; Jaeger et
al., 2010), and our timetree does not rule out a single colonization event for both platyrrhine
monkeys and caviomorph rodents to South America (Flynn and Wyss, 1998). e divergence
of New World and Old World primates has generally been estimated to either the Late Eocene
~37 Ma (Poux et al., 2006; Fabre et al., 2009) or the Middle Eocene ~43 Ma (Eizirik et al.,
2004; Janecka et al., 2007; Chatterjee et al., 2009). In their analysis of divergence times in both
monkeys and rodents, Poux et al. (2006) found evidence that caviomorphs diverged earlier in
the Eocene than platyrrhines, but could not exclude the possibility that both groups dispersed
contemporaneously to South America. If new fossil discoveries of South American primates can
establish a minimum age on the continent prior to ~27 Ma (Hostetter, 1969), it would sup-
port the case for concurrent trans-Atlantic dispersal with rodents (Bandoni de Oliveira et al.,
2009; Antoine et al., 2012). Earlier fossil nds for platyrrhine primates would also extend the
age of their crown radiation, which has been estimated to ~26 Ma (Opazo et al., 2006; Chat-
terjee et al., 2009) or ~15 Ma (Poux et al., 2006; Fabre et al., 2009), and close the substantial
gap relative to the ~40-30 Ma crown radiation of caviomorph rodents (Tab. 4).
The specter of paraphyly for caviomorphs (and rodents) has been raised on several
occasions, notably by studies that questioned if “guinea pigs are rodents” (e.g., Graur et al.,
1991; D’Erchia et al., 1996). ese early studies had woefully incomplete taxon sampling
Caviomorph evolution through time
and their reconstructions were plagued by long-branch attraction. One credible challenge is
that to Caviomorpha as a New World endemic, based on resemblances between the ~34 Ma
African fossil †Gaudeamus and possibly contemporaneous fossil caviomorphs from Santa Rosa,
Peru (e.g., †Eoincamys); however, ecological convergence presumably explains their dental
similarity (Sallam et al., 2009; Coster et al., 2010; Antoine et al., 2012). e polyphyletic
origin of caviomorphs involving two independent colonizations of South America has also
been proposed on the basis of cephalic arterial patterns (Bugge, 1971, 1985) and myology
(Woods and Hermanson, 1985). Incisor enamel microstructure shows similarities among South
American octodontoids and African thryonomyoids (Martin, 1994a; Martin, 2005), which
does not rule out polyphyly, but more parsimoniously suggests their common retention of an
ancestral condition. Recent studies condently support the monophyly of living caviomorphs
using multiple data sources, including cranial and dental characters (Marivaux et al., 2004;
MacPhee, 2011), DNA sequences (e.g., Nedbal et al., 1994; Meredith et al., 2011), and other
genomic elements (e.g., Huchon et al., 2007; Churakov et al., 2010).
Since Simpson (1945), there has been general agreement that living caviomorphs are divided
into four superfamilies, although with more debate over their content: Erethizontoidea with
a single family; Cavioidea with Caviidae, Dasyproctidae, Dinomyidae, and Hydrochoeridae;
Chinchilloidea with a single family; and Octodontoidea, with Abrocomidae, Capromyidae,
Ctenomyidae, Echimyidae, and Octodontidae (Patterson and Wood, 1982).e content of Octodon-
toidea was later expanded to include Myocastoridae (Woods, 1982), which had been treated as a
subfamily of Echimyidae. Although the traditional arrangement placed Dinomyidae in Cavioi-
dea, this family is now considered a chinchilloid (see also Spencer, 1987; Huchon and Douzery,
2001). For reasons mentioned above, African thryonomyids have also been placed in Octodon-
toidea (Simpson, 1945).
Relationships among these groups have slowly come into focus, limited mainly by inad-
equate taxonomic and genetic sampling. Using solely morphological evidence, McKenna and
Bell (1997) treated both Hystricidae and Erethizontidae as incertae sedis within Hystricognathi.
With a single gene, Huchon and Douzery (2001) recovered Erethizontoidea as sister to Octodon-
toidea + Chinchilloidea rather than Cavioidea. In an analysis that lacked chinchilloids, Vilela
et al. (2009) recovered erethizontiods as sister to cavioids + octodontoids. However, our tree
(Figs. 2 and 3) securely recovers the pairings Erethizontoidea + Cavioidea and Chinchilloidea +
Octodontoidea, which were also supported by Honeycutt (2009), Meredith et al. (2011), Fabre
et al. (2012a; 2012b), and Upham and Patterson (2012).
Living erethizontoids all belong to the New World porcupine family Erethizontidae,
which is recovered as strongly monophyletic. is family includes two lineages: Chaetomys on
the one hand and Erethizon and Coendou on the other, with Coendou now including species
previously allocated to Echinoprocta and Sphiggurus (Voss et al., 2013). Because it retains decidu-
ous premolars, Chaetomys had been considered a primitive echimyid (e.g., Patterson and Wood,
1982) until molecular data claried its relationships (Vilela et al., 2009). However, Martin
(1994b) had earlier correctly predicted the anity of Chaetomys with Erethizontidae based on
enamel microstructure. e lineage leading to Chaetomys diverged from other American por-
Upham and Patterson
cupines 17.5 Ma (Fig. 5), soon after Dinomyidae separated from Chinchillidae (19.7 Ma) and
Octodontidae separated from Ctenomyidae (18.9 Ma), and roughly coeval with the separation
of Capromyidae and Echimyidae (17.4-18.2 Ma) and of subfamilies within Caviidae (16.8-
17.6 Ma). is early divergence date and its morphological dierentiation justify distinguishing
Chaetomys in its own monotypic subfamily (cf. Martin, 1994b; Woods and Kilpatrick, 2005)
or maybe even family. e fact that all other South American porcupines diversied after their
divergence from Erethizon (7.5 Ma) also corroborates the decision of Voss (2011) to synonymize
Sphiggurus and Echinoprocta with Coendou.
Cavioidea is recovered as monophyletic, as are each of its families. Our analysis had moderate
support for Caviidae as sister to Cuniculidae + Dasyproctidae. Using an eight-gene supermatrix
and exemplar taxa, Fabre et al. (2012b) recovered Cuniculidae as sister to Dasyproctidae +
Caviidae (see also Huchon and Douzery, 2001). Using a combination of morphological and
molecular characters from four genes, Pérez and Pol (2012) posited a still dierent set of family
relationships: Dasyproctidae as sister to Cuniculidae + Caviidae. Surely, confusion over the
family-level relationships of cavioids is one of the major unresolved questions in caviomorph
systematics. Given the group’s rich fossil record, analyses that combine morphological and
molecular approaches (e.g., Pérez and Pol, 2012) are likely to be the most informative.
e Caviidae is currently thought to include three subfamilies: Dolichotinae, containing
Dolichotis; Hydrochoerinae, containing Hydrochoerus and Kerodon; and Caviinae, containing
Cavia, Microcavia, and Galea. Our reconstruction securely recovers Caviinae as sister to Hydro-
choerinae + Dolichotinae and Galea as sister to Cavia + Microcavia, agreeing in detail with the
analysis of Perez and Pol (2012). In addition, our timetree shows that the Hydrochoerinae-
Dolichotinae MRCA (13.2 Ma) postdates the divergence of Galea from other caviines (16.8
Ma; Fig. 5). Because this family has numerous fossil members not included in our analysis, and
because Fabre et al. (2012a) recovered a dierent topology for the Hydrochoerus clade, a com-
prehensive taxonomic revision is needed.
Chinchilloidea is strongly supported, as are its two families: Chinchillidae and Dinomyidae,
the latter represented by a single living species. Chinchillid relationships are securely recovered as
Lagostomus sister to Chinchilla + Lagidium. Spotorno et al. (2004) recovered the same topological
relationship in their analysis of cyt-b variation in species of Chinchillidae. Although some
have considered Abrocomidae as a member of this lineage (e.g., Glanz and Anderson, 1990),
abrocomids are now known to be members of Octodontoidea.
e extinct Miocene and Pliocene diversity of chinchilloids is impressive, with the extinct
families †Cephalomyidae and †Neoepiblemidae (Kramarz, 2002) and extinct dinomyid sub-
families †Eumegamyinae and †Potamarchinae (McKenna and Bell, 1997). e largest known
fossil rodent, †Josephoartigasia monesi, is a dinomyid from the late Pliocene of Uruguay with an
Caviomorph evolution through time
estimated body size of 350-1000 kg (Millien, 2008; Rinderknecht and Blanco, 2008). Clearly
there is a fuller understanding of chinchilloid evolution to be gained by considering the broader
geographic and ecological diversication of their fossil members in addition to their modern
(primarily Andean) representatives (Fig. 1.5).
Octodontoidea is strongly supported as monophyletic, with ve families: Abrocomidae,
Octodontidae, Ctenomyidae, Capromyidae, and Echimyidae (without Chaetomys but includ-
ing Myocastor). is content of the superfamily and families agrees with a developing consensus
among molecular phylogenetic studies (e.g., Huchon and Douzery, 2001; Leite and Patton, 2002;
Galewski et al., 2005; Fabre et al., 2012a; Fabre et al., 2012b). We securely recover Abrocomidae
as the sister lineage to the remaining octodontoids, a nding with important biogeographic im-
plications for the group as a whole (see Upham and Patterson, 2012). In an earlier single-gene
analysis of the vWF exon, Octodontidae appeared to be sister to all other lineages within the
superfamily (Huchon and Douzery, 2001). Again in agreement with traditional classications
(e.g., Patterson and Wood, 1982), we nd that the other octodontoid families form well-sup-
ported dyads: Octodontidae + Ctenomyidae and Capromyidae + Echimyidae.
Abrocoma and Cuscomys are the only living members of the family, so their robust
grouping in Fig. 3 is unsurprising. However, this is the rst molecular test of monophyly of this
family, executed here with complete ve-gene sampling and all plausible relatives. e modern
specimen of Cuscomys ashaninka we sequenced is the only one known from this genus; its congener
C. oblativa is known from historical material found alongside human burials at Machu Picchu
(Emmons, 1999). Abrocoma and Cuscomys diverged 8.4 Ma (Fig. 5), approximately coeval with
the split between the echimyid sister genera Lonchothrix and Mesomys.
As in earlier analyses of Octodontidae, there is strong evidence for monophyly and good
resolution of generic groupings. is group diverged from Ctenomyidae in the early Miocene
(Fig. 5), but crown-group octodontids did not split into the two main lineages until the late
Miocene, 8.8 Ma. One, the vizcacha-rat lineage, consists of a basal split of Octomys from the
group containing Pipanacoctomys as sister to Salinoctomys + Tympanoctomys. ese latter genera
are arid-adapted and appear to be closely related, sharing a MRCA 2.9 Ma, with the Salinoctomys-
Tympanoctomys divergence ~0.7 Ma during the mid-Pleistocene. Originally described by Mares
et al. (2000), Pipanacoctomys and Salinoctomys are considered in synonomy with Tympanoctomys
by some authors who question the morphological dierences among these taxa (Díaz and Verzi,
2006; Pérez, 2013). is issue is not resolved, but the genetic similarity of S. loschalchalerosorum
and T. barrerae indicates a divergence time younger than for any other caviomorph genus (Fig.
5).e recent description of a new species of vizcacha rat (T. kirchnerorum) (Teta et al., 2014),
and the putative whole genome duplication underlying their radiation (Suárez-Villota et al.,
2012), additionally call for a comprehensive revision of the vizcacha rats.
Upham and Patterson
e other octodontid lineage consists of Octodontomys as sister to Octodon (Spalacopus,
Aconaemys). Both Octodontomys and Octodon are terrestrial and/or saxicolous, whereas Spalacopus
+ Aconaemys appear adapted for fossoriality and, in the case of Spalacopus, subterranean life.
Spalacopus is characterized by strongly procumbent incisors and short ears and tail, all associated
with underground living (Lessa et al., 2008) and clearly derived conditions within octodontids.
e divergence of Spalacopus and 2 species of Aconaemys (fuscus and sagei) is dated as 3.1 Ma, in the
late Pliocene; however, Aconaemys was found to be paraphyletic in our species-level analyses (not
shown), with A. porteri weakly supported as sister to the remaining taxa. e paraphyly of Spalacopus
and Aconaemys has been documented elsewhere (Gallardo and Kirsch, 2001; Honeycutt et al.,
2003; Upham and Patterson, 2012) and calls for a revision of this group, particularly considering
the similarity between Aconaemys and the Plio-Pleistocene form †Pithanotomys (Reig, 1986).
Our analysis recovered Ctenomys, the sole extant genus of ctenomyids, as a strongly sup-
ported monophyletic lineage that diverged from the Octodontidae roughly 18.9 Ma. However,
most early branches in this family (all genera except Ctenomys) went extinct, and †Ctenomys
uquiensis places a minimum age on the genus of ~3.5 Ma (Verzi et al., 2010). We dated the
crown Ctenomyidae radiation at 6.0 Ma (4.6-7.6), which diers from three estimates based ex-
clusively on cyt-b data: 5.1 Ma (3.3-6.9; Lessa and Cook, 1998), 3.7 Ma (Castillo et al., 2005),
and 9.2 Ma (6.4-12.6; Parada et al., 2011). Including data from two low-variability introns,
Castillo et al. (2005) also estimated an age of 1.3 Ma that they recognized was inconsistent with
the fossil record. Given the minimum inferred divergences of Ctenomys-†Praectenomys ~4 Ma
and crown Ctenomyidae ~6.5 Ma (Verzi et al., 2013), the fossil record agrees with molecular
estimates that place the crown tuco-tuco radiation in the latest Miocene or earliest Pliocene.
e Capromyidae (hutias) are recovered as monophyletic and well resolved, although
they are nested within Echimyidae. e Hispaniolan Plagiodontia appears as the basal member
of this clade, with the Jamaican and Bahamian Geocapromys as the next-most basal. ese genera
ank a clade of exclusively Cuban hutias, with Capromys as sister to Mysateles + Mesocapromys.
e divergence of the three Cuban genera dates only to 3.0 Ma, in the Late Pliocene. In fact,
Mesocapromys and Mysateles were regarded as subgenera of Capromys until Kratochvil et al.
(1978) elevated them to generic rank (but see Woods, 1989). In view of their recent origins
and morphological and genetic distinctions (e.g., Camacho Pérez et al., 1995; Borroto-Páez
and Mancina, 2011; Kilpatrick et al., 2012), this group also requires a revision with combined
molecular and cladistic character analysis of fossil and living taxa.
e mid-Miocene (17.4 Ma) split between Capromyidae and a lineage of Brazilian echimyids
considerably postdates GAAR (Greater Antilles + Aves Ridge; Fig. 6.2), a transitory corridor
for dispersal from northern South America to the Caribbean ~34 Ma (Iturralde-Vinent and
MacPhee, 1999).e delayed 9.8 Ma age for crown Capromyidae supports the idea that a
considerable portion of this radiation is now extinct (Woods et al., 2001; Borroto-Páez and
Mancina, 2011). e oldest capromyid fossil, †Zazamys, was here assigned to the Capromyidae
Caviomorph evolution through time
crown group as a stem member of Isolobodontinae following MacPhee et al. (2003), but this
calibration point was excluded from the nal analyses due to concerns about the signicantly
older node age upon its inclusion (Fig. 4.2). Until the phylogenetic position of †Zazamys relative
to modern capromyids and recently extinct taxa (App. 1) can be condently established, using
this fossil as a crown constraint is questionable (see also Fabre et al. 2014).
Our phylogenies identify three lineages of spiny rats (numbered “I”, “II”, and “III” in
Figs. 3 and 4), two of which constitute well-supported clades. However, none support the tra-
ditional subdivision of living echimyids into the subfamilies Echimyinae, Dactylomyinae, and
Eumysopinae (e.g., Woods and Kilpatrick, 2005). Instead, echimyines appear in both lineages
I and II, dactylomyines comprise a small part of a lineage I, and eumysopines are scattered
across all three clades. e lineages we recover also conict with the latest cladistic analyses of
morphological variation in fossil and living taxa (e.g., Olivares et al., 2012). Nevertheless, our
results agree with previous molecular studies in areas of overlap and strong statistical support
(e.g., Leite, 2003; Galewski et al., 2005).
We recovered strong support for an arboreal lineage (I) that comprises all dactylomyines,
most echimyines, plus a few eumysopines. is group includes the genera Lonchothrix, Mesomys,
Isothrix, Santamartamys, Diplomys, Olallamys, Kannabateomys, Dactylomys, Pattonomys, Toromys,
Makalata, Echimys, and Phyllomys. Some of these genera belong to well-supported dyads:
Lonchothrix + Mesomys, Santamartamys + Diplomys, Pattonomys + Toromys, and Echimys +
Phyllomys. Others, including Isothrix and Makalata, have less certain placements within the
group. e three bamboo rat genera (Olallamys, Kannabateomys, and Dactylomys) are strongly
supported as a group that shared a MRCA 10.3 Ma, but within that clade are unresolved, with
weak support for a sister-group relationship between Kannabateomys and Dactylomys. Given
the morphological and ecological distinctiveness of bamboo rats, the recovery of two erstwhile
echimyine genera, Diplomys and Santamartamys, as their sister clade is remarkable. is is
the rst time Olallamys, Diplomys, and Santamartamys have been included in any molecular
phylogeny, expanding the membership of this clade to non-bamboo rats and centering its
geographic distribution in the Northern Andes.
We also found strong support for a widespread and ecologically diverse lineage (II), of which
the terrestrial rodent richomys appears to be the basal member. Our timetree (Fig. 5) dates
the stem divergence leading to richomys as 15.5 Ma. Using extensive sampling of richomys
populations across their geographic range, Nascimento et al. (2013) dated the richomys crown
group to the Late Miocene (~8.5 Ma). e next-most-basal member of this clade is Myocastor,
which was thought to represent a separate family, Myocastoridae (Woods, 1993), until molecu-
lar phylogenetic analyses consistently recovered it as an echimyid (e.g., Leite and Patton, 2002).
Myocastor is securely recovered as sister to a triad that includes the eumysopine pair Proechimys
+ Hoplomys plus the echimyine Callistomys. For most of the time since its initial description
(Pictet, 1841), Callistomys pictus was recognized as a species of Echimys or Isothrix, both
echimyines. Emmons and Vucetich (1998) recognized many dierences in cranial characters
between this form and other living echimyids and erected a new genus for it and a related
fossil form. Recovery of Callistomys in lineage II with semi-aquatic Myocastor and the terrestrial
Upham and Patterson
spiny rats Proechimys + Hoplomys suggests that its soft uy pelage and arboreal or scansorial
habits evolved independently from similar traits in lineage I (see also Upham, 2014; Loss et al.,
2014). Resemblances between Callistomys and extinct echimyid lineages in the mid-Miocene
and earlier (e.g., †Maruchito; Emmons and Vucetich, 1998; Verzi et al., 2013) make it likely that
the nearest relatives of Callistomys are missing from our phylogenetic tree.
e third echimyid lineage (III) is only partially supported and creates topological and no-
menclatural issues for the Echimyidae by rendering the family paraphyletic with respect to
Capromyidae. Galewski et al. (2005) were rst to identify a clade of three Brazilian Shield taxa
–Trinomys as sister to Clyomys + Euryzygomatomys– that joined to Capromys rather than the re-
maining echimyids (see also Fabre et al., 2012b; Upham and Patterson, 2012).is relationship
is unchanged by the addition of four additional capromyid genera, which join Capromys to form
a robustly monophyletic group. It is also unaltered by the inclusion of Carterodon, a Cerrado
endemic, which is recovered as sister to the other Brazilian Shield taxa or falls into a trichotomy
with them and the capromyids (Fig. 3). Whereas the fossorial Clyomys and Euryzygomatomys
diverged in late Miocene times (5.7 Ma; Fig. 5), other members of this lineage have much
longer branches. e lineage leading to Trinomys, which was considered merely a subgenus of
Proechimys until the molecular study of Lara et al. (1996), diverged from other echimyids 15.8
Ma, and the branch to Carterodon diverged still earlier, at 16.7 Ma. Carvalho and Salles (2004)
grouped Carterodon, Clyomys, and Euryzygomatomys together with some fossil forms at the base
of crown-group Echimyidae, which is consistent with our results. Olivares et al. (2012) found
support for the Miocene fossil †eridomysops as sister to Clyomys + Euryzygomatomys, joined
next as sister by Carterodon, but they recovered Trinomys as sister to Proechimys rather than a
member of this clade. Resolving this node may be clouded by the extinction of †eridomysops
and other forms, which would have pruned relatives that might help to resolve it.
ere is also the possibility of a hard polytomy at the base of some echimyid lineages stemming
from rapid or simultaneous divergence, an idea rst posed to explain the lack of resolution
in mtDNA phylogenies (Lara et al., 1996; Leite and Patton, 2002). e inclusion of slower-
evolving nuclear exons helped resolve some elements of this “star-phylogeny” (e.g., Galewski et
al., 2005; Fabre et al., 2012b), but it did not exclude the possibility that unresolved nodes were
real polytomies (hard) rather than being due to lack of data (soft). Our expanded molecular
sampling of echimyid genera helps to solidify the sister relationship between lineages I and II,
and resolve the position of several Miocene-aged nodes (Figs. 3 and 5). Group membership
for previously unsampled taxa (e.g., Pattonomys, Santamartamys) is also claried. Nevertheless,
key nodes remain unresolved in lineage I (relative placement of Isothrix, Mesomys /Lonchothrix,
bamboo rats, and tree rats) and, as detailed above, Carterodon relative to the clades of Antillean
hutias and Brazilian echimyids. Not to be ignored, however, is the potential resolving power
that molecular sequences from missing echimyids may still have on their inferred phylogeny.
In particular, the historical extinction of Caribbean echimyids allied to †Heteropsomyinae
(e.g., Boromys; App. 1) presents a fascinating opportunity to include ancient DNA from this
lineage. Considering echimyid fossils in this assessment is also necessary. Several mid-Miocene
echimyids (e.g., †Stichomys) cluster with living members of lineage I in recent cladistic analyses
(Verzi et al., 2013; Arnal et al., 2014), reinforcing the idea that resolving the early history of
Echimyidae hinges upon combined analyses of morphological and molecular diversity.
Caviomorph evolution through time
Environmental context for caviomorph rodent diversication
Geological and paleoclimatic studies allow us to glimpse the varied and dynamic envi-
ronments that caviomorph ancestors would have experienced during their Middle Eocene-Re-
cent evolutionary history (Fig. 6). Caviomorph ancestors inhabited a hot and wet “greenhouse”
world that lacked polar icecaps and had mean global temperatures >10 °C warmer than they are
today (Zachos et al., 2008; Goin et al., 2012). A dramatic global cooling event unfolded near
the Eocene-Oligocene boundary that is linked to the opening of the Drake Passage between
Cape Horn and Antarctica, formation of a strong Antarctic Circumpolar Current, and ensuing
Antarctic glaciation (Livermore et al., 2004; Zachos et al., 2008; Lagabrielle et al., 2009).
Figure 6. The timing of 1. divergence events throughout the molecular timetree of Ctenohystrica (grouped in 1 Ma bins from
the species-level timetree [not shown]) in relation to 2. contemporaneous climatic and geologic events. Note that only the
surviving lineages are represented in the timetree. The oxygen isotopic curve is modied from Zachos et al. (2008) and is used
to represent global climatic changes. The temperature scale assumes an ice-free ocean, so only applies directly to the period
prior to ~35 Ma (Eocene–Oligocene boundary), after which a global cooling event initiated ice sheets in Antarctica and a
shift to an “icehouse” world (Livermore et al., 2004; Lagabrielle et al., 2009; Goin et al., 2012). Other references for geoclimatic
events are as follows: the GAAR (Great Antilles/Aves Ridge) land connection ~34 Ma for dispersal to the Caribbean from
South America (Iturralde Vinent and MacPhee, 1999); global drop in sea levels ~11 Ma (Haq et al., 1987); orogenic episodes
and percentages of modern elevations in the Central Andes 22-20 Ma and 12-10 Ma, and Northern Andes 5-2 Ma (Gregory–
Wodzicki, 2000; Lomize, 2008); Altiplano-Puna rainshadow by 15 Ma (Hartley, 2003); Pebas wetlands in Amazonia 23-11 Ma
and terra rma forests beginning ~7 Ma (Hoorn et al., 2010; Latrubesse et al., 2010; Shephard et al., 2010); establishment of
the east owing Amazon River between 10.6 and 9.7 Ma (Figueiredo et al., 2010); and stages of decreasing forest cover in
Patagonia (Barreda and Palazzesi, 2007; Barreda et al., 2010). The timing of geological epochs is from Gradstein et al. (2004).
Upham and Patterson
e subsequent “icehouse” world provided the backdrop for Caviomorpha’s initial diversi-
cation ~32 Ma into four strongly dierentiated superfamilies (Figs. 5 and 6). Several stages
of Andean orogeny during the Miocene are well supported, with the rst major uplift in the
Central Andes by 20 Ma, followed by a later uplift 12-10 Ma to half of its modern elevation of
~3700 m in the Altiplano-Puna (Gregory-Wodzicki, 2000; Lomize, 2008). By ~15 Ma, Andean
uplift created a rainshadow across much of eastern South America, especially in the south (Hartley,
2003), leading to a striking process of aridication throughout Patagonia and the Southern
Cone. e middle to late Miocene drying of Patagonia reduced a formerly wet, forested biome
to arid shrubland, desert, and relict montane forests (Barreda and Palazzesi, 2007; Barreda et
al., 2010). Major growth of the Northern Andes was not triggered until ~5 Ma, but subsequent
uplift was rapid. By 2 Ma, the full modern elevation of both the Central and Northern Andes
had been reached (Gregory-Wodzicki, 2000). Meanwhile in the Amazon Basin, the early stages
of Andean orogeny took place as Western Amazonia was covered by the inland Pebas system
of wetlands and oodplains, which lasted from ~23-11 Ma (Hoorn et al., 2010; Latrubesse et
al., 2010). Shifting drainage patterns to the east and lowered global sea levels led to the Pebas
system receding and establishing the east-owing Amazon River (Haq et al., 1987; Figueiredo
et al., 2010), so that by ~7 Ma, terra rma rainforests had expanded widely in the Amazonian
lowlands (Hoorn et al., 2010; Shephard et al., 2010; Bonvicino and Weksler, 2012).
ese geoclimatic events –individually and in concert– created major changes in South
American biomes that certainly would have impacted the evolutionary diversication of
caviomorphs. For example, episodes of Andean uplift provided novel highland habitats and
opportunities for genetic isolation, promoting novel variants that might subsequently recolo-
nize lowland habitats as incipient species. Evidence for transitions between lowland and high-
land habitats has been amassed within clades of Echimyidae (bamboo rats, spiny tree-rats, and
brush-tailed rats; Upham et al., 2013), as well as Andes-Amazon lineages of frogs (e.g., Santos
et al., 2009), birds (e.g., Weir, 2006), and mammals such as opossums (de la Sancha et al.,
2012) and olingos (Helgen et al., 2013). Both directions of transition (lowland-to-highland and
highland-to-lowland) appear to have been common in South America since the late Miocene
(Upham et al., 2013), supporting the thesis that processes of geoclimatic and biological evolu-
tion are intricately linked (Patterson et al., 2012; Morrone, 2014).
Perhaps most dramatically for caviomorph rodents was the major dierentiation of South
American habitats into arid southern and mesic northern biomes after ~18 Ma. Substantial
global cooling following the Mid-Miocene Climatic Optimum, along with the initiation of a
rainshadow in most of the Southern Cone (Fig. 6.2), transformed the continent into multiple
distinctive biomes, each with unique environmental challenges to resident caviomorphs (Croft
et al., 2009; Flynn et al., 2012). e fossil record of Octodontoidea appears to especially reect
these changes, with the mesic-adapted clade of Echimyidae-Capromyidae represented in Pata-
gonian localities during the Miocene (e.g., Vucetich et al., 1993), but replaced thereafter by a
greater diversity of arid-adapted forms allied to Octodontidae-Ctenomyidae (Verzi et al., 2008;
Verzi et al., 2013). According to our molecular timetree, these main clades of Octodontoidea
have approximately coeval crown diversications (18.2 Ma and 18.9 Ma, respectively; Fig. 5),
roughly concordant with the timing of continent-wide climatic dierentiation. is poses a
key question about the subsequent evolution of these clades: What factors led to the greater
Caviomorph evolution through time
longevity of lineages in Echimyidae-Capromyidae (many Miocene-aged lineages) compared to
those in Octodontidae-Ctenomyidae (two surviving lineages from the late Miocene)? e fact
that the southern clade of octodontids and ctenomyids had to adapt to arid climates unprec-
edented in the Paleogene (Fig. 6.2), suggests one reason for the apparently greater turnover of
species during the initial stages of their radiation (Fig. 5; Upham, 2014). Spatiotemporal and
ecomorphological data from the fossil record need to be reconciled with the region’s progressive
aridication, especially now that rates and sequences of lineage diversication can be inferred
from the molecular timetree.
e juxtaposition of caviomorph diversication events (Fig. 6.1) along with the climatic and
environmental factors that might have shaped their evolutionary history (Fig. 6.2) is intended
only to be heuristic; we emphasize it is not exhaustive. In another study, we examine the tempo
and mode of caviomorph diversication and disparication more directly. However, Figure 6
oers a glimpse of the major environmental factors that would have shaped caviomorph evolution
from the Oligocene-Miocene, prior to the arrival of other rodent groups (including squirrels,
beavers, and gophers) and placental lineages to South American ecosystems (Simpson, 1980).
e opportunities that caviomorphs encountered and were able to exploit are directly reected
in their diversity and disparity, but we are still far from understanding this complementarity.
e remarkable fossil and living diversity of caviomorph rodents deserves to be highlighted
in evolution textbooks – this is truly an exemplar lineage for studying biological diversity and
diversication through time.
Sincere thanks are due to the organizers A. Vassallo and D. Antenucci for their eorts
organizing this volume. For access to tissues, we thank the following individuals and institutions:
R. Borroto-Páez; R. Voss (AMNH); B. Stanley and S. Goodman (FMNH); J. Patton and C.
Conroy (MVZ); M. Mares and J. Braun (SNOMNH); Y. Leite, L. Costa, and C. Loss (UFES);
L. Emmons (USNM); E. Sargis (YPM); B. Kilpatrick (ZTNH); A.P. Carmignotto, R. Paresque,
and R. Moura. For access to laboratories and assistance with molecular work, we thank: K.
Feldheim and B. Wray (FMNH); B. Lim, K. Choe, and J. Miller (ROM); H. Poinar, M.
Kuch, and J. Enk (MADC); and Y. Leite, C. Loss, and R. Guimarães (UFES). E. Lessa and J.
Patton oered insightful comments throughout our analyses. We also want to extend thanks to S.
Lidgard for discussions ultimately leading to Fig. 6; to D. Jablonski, T. Price, and R. Ree for helpful
comments as members of N.S.U.’s dissertation committee; to H. Mantilla-Meluk for generous
help with castellano; and to N. Stewart for outstanding editorial support throughout the course
of this study. is research was conducted under an NSF Doctoral Dissertation Improvement
Grant (DEB-1110805), ASM Grant-In-Aid of Research, Hinds Fund Grant from the University
of Chicago, and funds from the FMNH Pritzker Laboratory for Molecular Systematics.
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Caviomorph evolution through time
Appendix 1. Taxonomy and genetic sampling for this study, showing GenBank accession numbers for the ve gene regions examined. Sequences in bold were generated newly
for this study or Upham et al. (2013), and are associated with the listed museum voucher or collector numbers. Species listed without reference numbers are entirely from GenBank
and are specimen chimeras in most cases. Species denoted with * lacked published DNA sequences prior to this study, and those with ‡ were sequenced from dried tissue. “NS”
accompanies species not sampled here, and † denotes those recently extinct. This listing is updated from Woods and Kilpatrick (2005) using the IUCN Mammal Redlist (2012) and
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
Pedetes species chimerabAJ389527 (s) AY012113 (c) AF332025 (c) AJ238389 (s) AY011882 (c)
Laonastes aenigmamus AM407933 DQ139934 AM407901 AM407897
Ctenodactylus species chimeracAJ389532 (v) AJ389543 (v) AF332042 (g) JN415077 (g) JN633629 (g)
Felovia vae NS
DĂƐƐŽƵƟĞƌĂŵǌĂďŝ AJ389533 AJ389544
Atherurus africanus HQ450774 AY093658
Atherurus macrourus FJ931121 U12451 JF938865 AJ251131
Hystrix africaeaustralis X70674 U12448 AF332033
Hystrix brachyura JQ991599 AY012117 JN414760 JN415082 AY011886
Hystrix crassispinis NS
Hystrix cristata FJ472565 AY093659
Hystrix indica AY692229 AY093669
Hystrix javanica NS
Hystrix pumila NS
Hystrix sumatrae NS
Trichys fasciculata FM162081 AJ224675
ĂƚŚǇĞƌŐƵƐũĂŶĞƩĂ AF012241 AY425843
Bathyergus suillus AY425913 FM162080 AJ238384
Fukomys amatus AF012233 AY427021
Fukomys anselli AY427022
Fukomys bocagei AF012229 AF012213
Fukomys damarensis AF012220 AY427026 FN984748 FN984751
Upham and Patterson
Fukomys darlingi AF012232 AY427033
Fukomys foxi AY427036
Fukomys ilariae NS
Fukomys kafuensis AY427037
Fukomys mechowi AF012230 AY427041
Fukomys ochraceocinereus AY427045
Fukomys whytei AY425863 AY427046
Fukomys zechi NS
ƌǇƉƚŽŵǇƐŚŽƩĞŶƚŽƚƵƐ AY425885 AY427064 FJ855202 AJ251132
Cryptomys anomalous AY427054
Cryptomys holosericeus AY427051
Cryptomys natalensis M63568
Cryptomys nimrodi NS
Georychus capensis AF012243 AY427066 FJ855203
Heliophobius argenteocinereus FMNH 168421 KJ742646 AY427070 FJ855204 AJ251133 KJ742671
Heterocephalus glaber AF155870 AY427075 AF332034 AJ251134 AY011889
Petromus typicus DQ139935 M63571 JN414761 AJ251144 JN633636
Thryonomys gregorianus NS
Thryonomys swinderianus FMNH 165618 KJ742647 NC_002658 AF332035 AJ224674 KJ742672
Chaetomys subspinosus EU544660
Erethizon dorsatum KC463889 AY012118 AF332037 AJ251135 AY011887
Coendou bicolor KC463860
Coendou ichillus KC463861
Coendou insidiosus JX312693 JX312693
Coendou melanurus KC463862 AJ389549 AJ224664
Coendou mexicanus KC463863 FJ855212
Coendou nycthemera KC463864
Coendou prehensilis ĐŽƌƌĞĐƟŽŶdKC463873 AF520695 AF520663
Coendou pruinosus KC463880
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
Caviomorph evolution through time
Coendou quichua KC463882
Coendou roosmalenorum NS
Coendou rufescens KC463884
Coendou spinosus KC463887
Coendou villosus NS
Cavia aperea GU136753 AF433908 AF433930
Cavia fulgida GU136737
Cavia intermedia NS
Cavia magna GU136734 AY765986
Cavia porcellus AF490405 AF433909 AF433931 AJ224663 XM_003463833
Cavia tschudii GU136731 AY012121 FJ855206 AY011890
Galea musteloides FMNH 164943 KJ742648 AF433910 AF433933 KJ742608 KJ742673
Galea spixii GU067491 AF433913 AF433935
Microcavia australis AF491750 AF433915 AF433937
Microcavia niata GU136725
Microcavia shiptoni NS
ŽůŝĐŚŽƟƐƉĂƚĂŐŽŶƵŵ GU136724 AF433917 AF433939
ŽůŝĐŚŽƟƐƐĂůŝŶŝĐŽůĂ GU136723 AF433919 AF433940
Hydrochoerus hydrochaeris GU136721 AF433924 FJ855208 AJ251137 AY011891
Hydrochoerus isthmius NS
Kerodon acrobata GU477346
Kerodon rupestris GU136722 AY765988 AF433938
Dasyprocta azarae NS
Dasyprocta coibae NS
Dasyprocta cristata NS
Dasyprocta fuliginosa AF437784
Dasyprocta guamara NS
Dasyprocta kalinowskii NS
Dasyprocta leporina AF437791 AY093660 FJ855207 U31607
Dasyprocta mexicana NS
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
Upham and Patterson
Dasyprocta prymnolopha NS
Dasyprocta punctata AF433921 AF433943 JN415079
Dasyprocta ruatanica NS
Myoprocta acouchy FMNH 160003 KJ742649 AF433922 AF433945 KJ742609 KJ742695
DǇŽƉƌŽĐƚĂƉƌĂƫ U34850 AF433923 AF433946
Cuniculus paca AY206555 AF520693 AF433928 AJ251136
Cuniculus taczanowskii FMNH 170721 KJ742656 AY012125 AF433929 JN415074 AY011894
Chinchilla chinchilla NS
Chinchilla lanigera FMNH 178049 AF464760 AF520696 AF332036 AJ238385 KF590658
Lagidium peruanum AY254885
Lagidium viscacia AY254886 FJ855209
Lagostomus crassus NS
Lagostomus maximus AF245485 FJ855210
Dinomys branickii AY254884 AY012124 AF520659 AJ251145 AY011893
ďƌŽĐŽŵĂďĞŶŶĞƫŝ AF244387 FJ855213 AJ251143 JN633625
Abrocoma boliviensis*‡ MVZ 120238 KJ742657
Abrocoma budini NS
Abrocoma cinerea AF244388 AF520666 AF520643
Abrocoma schistacea NS
Abrocoma uspallata NS
Abrocoma vaccarum NS
Cuscomys ashaninka * LHE 1359 KJ742658 KJ742598 KJ742626 KJ742610 KJ742683
†ƵƐĐŽŵǇƐŽďůĂƟǀƵƐ NS e
Aconaemys fuscus AF405351 AF520674 AF520657
Aconaemys porteri AF520671 AF520644
Aconaemys sagei MVZ 163419 KJ742650 AF520672 AF520645 KJ742675
Octodon bridgesi MVZ 184958 KJ742651 AF520676 AF520646 KJ742611 KJ742676
Octodon degus AF007058 AF520678 AF520647
Octodon lunatus AF227514 AF520681 AF520650 AJ238386
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
Caviomorph evolution through time
Octodontomys gliroides FMNH 162890 AF370706 AF520683 AF520649 KF590672 KF590663
Octomys mimax GQ121097 AF520686 AF520652
Pipanacoctomys aureus GQ121117 AY249753 AY249752
Salinoctomys loschalchalerosorum* CML 3695 KJ742652 KJ742607 KJ742635 KJ742612 KJ742684
Spalacopus cyanus AF007061 AF520688 AF520653
Tympanoctomys barrerae AF007060 AF520691 AF520655
Ctenomys australis AF370697
Ctenomys azarae JN791407
Ctenomys bergi AF144284
Ctenomys boliviensis AF007038 U12446 FJ855214 JN633630
Ctenomys brasiliensis NS
Ctenomys budini NS
Ctenomys colburni HM777474
Ctenomys coludo NS
Ctenomys conoveri AF007054
Ctenomys coyhaiquensis FMNH 134300 AF119112 KF590700 KF590678 KF590666 KF590659
Ctenomys dorbignyi AF500044
Ctenomys dorsalis NS
Ctenomys emilianus NS
Ctenomys famosus NS
Ctenomys fochi NS
Ctenomys fodax HM777475
Ctenomys frater AF007045
Ctenomys fulvus AF370688
Ctenomys goodfellowi AF007050
Ctenomys haigi AF422920 AF422853
Ctenomys ibicuiensis new species fJQ389020
Ctenomys johannis NS
Ctenomys juris AF144275
Ctenomys lami HM777477
Ctenomys latro HM777478
Ctenomys leucodon AF007056 HM544131
Ctenomys lewisi AF007049
Ctenomys magellanicus HM777479
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
Upham and Patterson
Ctenomys maulinus AF370703 AJ251138
Ctenomys mendocinus HM777480
Ctenomys minutus HM777483
Ctenomys occultus HM777485
Ctenomys opimus AF370700
Ctenomys osvaldoreigi NS
Ctenomys pearsoni HM777486
Ctenomys perrensi HM777489
Ctenomys peruanus NS
Ctenomys pilarensis AF144265
Ctenomys porteousi AF370682
Ctenomys pundit HM777490
Ctenomys rionegrensis AF119103 HM544130
Ctenomys roigi HM777492
Ctenomys saltarius HM777493
Ctenomys scagliai HM777494
Ctenomys sericeus HM777496
Ctenomys sociabilis HM777495 HM544129
Ctenomys steinbachi AF007044 AF520667 AF520656
Ctenomys talarum AF370699
Ctenomys torquatus AF119111
Ctenomys tuconax AF370684
Ctenomys tucumanus AF370691
Ctenomys tulduco NS
Ctenomys validus NS
Ctenomys viperinus NS
Ctenomys yolandae AF144285
Dactylomys boliviensis L23339 AF422875 JX515334 AJ849307 EU313298
Dactylomys dactylinus USNM 579620 L23335 AF422874 KF590681 KF590667 EU313300
Dactylomys peruanus EU313206
Kannabateomys amblyonyx AF422917 AF422850 AJ849310
Olallamys albicauda *‡ FMNH 71128 KF590697 KF590690 KF590673
Olallamys edax NS
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
Caviomorph evolution through time
Callistomys pictus* RM 233 KJ742659 KJ742594 KJ742627 KJ742614 KJ742677
Diplomys caniceps NS
Diplomys labilis*‡ FMNH 70101 KJ742660 KJ742636 KJ742613 KJ742685
Santamartamys rufodorsalis*‡ AMNH 34392 KJ742664
Echimys chrysurus L23341 AF422877 JX515333 AJ251141 EU313303
Echimys saturnus NS g
Echimys vieirai NS
WĂƩŽŶŽŵǇƐŽĐĐĂƐŝƵƐΎ‡ FMNH 84259 KJ742661 KJ742637
WĂƩŽŶŽŵǇƐƐĞŵŝǀŝůůŽƐƵƐΎ‡ FMNH 69118 KJ742662 KJ742616 KJ742687
Isothrix barbarabrownae ‡ FMNH 170722 EU313214 KF590701 KF590682 KF590668 EU313304
Isothrix bistriata L23349 JX515336 AJ849308 EU313307
Isothrix negrensis mislabeled hL23355 AF422873
Isothrix orinoci‡ USNM 406370 EU313223 KF590702 KF590683 KF590669 KF590660
Isothrix pagurus ‡ USNM 555639 EU313227 KF590703 KF590684 KF590670 KF590661
Isothrix sinnamariensis ROM 106624 AY745734 KF590704 KF590685 AJ849309 EU313312
Toromys grandis ‡ FMNH 92198 KF590699 KF590694 KF590676 EU313336
Toromys rhipidurus *‡ FMNH 87244 KJ742663 KJ742638 KJ742617 KJ742686
Makalata didelphoides UFMG 3012 L23362 KJ742600 KJ742639 AJ849311 KJ742688
Makalata macrura MVZ 153637 iL23356 AF422879 KF590687 AJ849312 EU313328
Makalata obscura NS j
Phyllomys blainvillii MVZ 197568 JF297836 KF590706 KF590692 JF297734 KF590664
Phyllomys brasiliensis EF608182 JF297729
Phyllomys dasythrix MCNU 844 JF297832 KJ742605 KJ742641 JF297708 KJ742689
Phyllomys kerri NS
Phyllomys lamarum EF608181 JF297730
Phyllomys lundi EF608183 JF297721
WŚǇůůŽŵǇƐŵĂŶƟƋƵĞŝƌĞŶƐŝƐ EF608179 JF297720
Phyllomys medius NS
Phyllomys nigrispinus JF297807 JF297714
WŚǇůůŽŵǇƐƉĂƩŽŶŝ MN 62391 EF608187 KJ742606 KJ742642 JF297744 KJ742690
Phyllomys sulinus JF297833 JF297710
Phyllomys thomasi NS
Phyllomys unicolor NSk
Carterodon sulcidens* LGA 735 KJ742666 KJ742596 KJ742640 KJ742615 KJ742678
Clyomys bishopi NS
ůǇŽŵǇƐůĂƟĐĞƉƐ MCNM 2009 AF422918 KJ742597 KJ742628 AJ849306 KJ742679
Euryzygomatomys spinosus UFMG 1948 EU544667 AF422854 KJ742629 AJ849319 KJ742680
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
Upham and Patterson
Hoplomys gymnurus AF422922 AF520668 AF520661 JN415080 JN633632
Lonchothrix emiliae AF422921 AF422857
Mesomys hispidus MEPN 12212 KF590705 KF590696 KF590688 KF590671 KF590662
Mesomys occultus MVZ 194396 L23388 AF422858 KF590689 EU313331
DĞƐŽŵǇƐƐƟŵƵůĂǆ UFROM 379 KJ742667 KJ742603 KJ742630 KJ742618 KJ742674
Myocastor coypus EU544663 AF520669 AF520662 AJ251140 AY011892
Proechimys brevicauda NS
Proechimys canicollis NS
Proechimys chrysaeolus NS
Proechimys cuvieri FMNH 175256 AJ251400 KF590707 KF590693 KF590675 KF590665
Proechimys decumanus NS
Proechimys echinothrix NS
Proechimys gardneri NS
Proechimys goeldii NS
Proechimys guairae NS
Proechimys guyannensis AJ251396
Proechimys hoplomyoides NS
Proechimys kulinae NS
Proechimys longicaudatus MVZ 197574 HM544128 HM544128 KJ742643 KJ742619 KJ742681
Proechimys magdalenae NS
Proechimys mincae NS
Proechimys oconnelli NS
Proechimys poliopus NS
Proechimys quadruplicatus U35413 AF422863 AJ849313
WƌŽĞĐŚŝŵǇƐƌŽďĞƌƟ EU544666 AJ251139
Proechimys semispinosus NS
Proechimys simonsi FMNH 175283 U35414 AF422864 KJ742631 AJ849320 EU313332
Proechimys steerei NS
Proechimys trinitatus NS
Proechimys urichi NS
Thrichomys apereoides EU313252 AF422855 JX515325 AJ849315 EU313334
Thrichomys inermis AY083343
Thrichomys pachyurus AY083329
Trinomys albispinus U34856
Trinomys dimidiatus UFMG 1951 U35169 AF422867 KJ742620 KJ742682
Trinomys eliasi U35166 AF422869
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
Caviomorph evolution through time
Trinomys iheringi FMNH 141667 EU313254 AF422868 KF590695 KF590677 EU313337
Trinomys mirapitanga NS
Trinomys moojeni NS
Trinomys myosuros NS
Trinomys paratus U35165 AF422866 AJ849316
Trinomys setosus AF422924 AF422871 AJ849317
Trinomys yonenagae AF194295 AF422865 AJ849318
†Boromys torrei NS
†Brotomys contractus NS
†Brotomys voratus NS
†Heteropsomys insulans NS
Capromys pilorides AF422915 AF433926 AF433950 AJ251142 JN633628
Geocapromys browni* ZTNH844 KJ742653 KJ742599 KJ742644 KJ742621 KJ742692
Geocapromys ingrahami*‡ FMNH 5624 KJ742668
†Geocapromys columbianus NS
†Geocapromys thoracatus NS
Mesocapromys angelcabrerai* RBP-A41 KJ742654 KJ742595 KJ742632 KJ742622 KJ742694
Mesocapromys auritus* RBP-B15 KJ742655 KJ742601 KJ742633 KJ742623 KJ742693
Mesocapromys melanurus* RBP-A25 KJ742669 KJ742602 KJ742691
Mesocapromys nanus NS
Mesocapromys sanfelipensis NS
Mysateles garridoi NS
Mysateles gundlachi NS
Mysateles meridionalis NS
Mysateles prehensilis * RBP-B23 KJ742670 KJ742604 KJ742634 KJ742624 KJ742696
WůĂŐŝŽĚŽŶƟĂĂĞĚŝƵŵ‡ FMNH 63876 KJ742665 KJ742645
ZTNH 843 KJ742625 KJ742697
†Hexolobodon phenax NS
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
Upham and Patterson
ĐǇƚͲbϭϮ^ƌZE ',Z ǀt& Z'ϭ
†Isolobodon montanus NS
†Isolobodon portoricensis NS
†Clidomys osborni NS
†Amblyrhiza inundata NS l
†Elasmodontomys obliquus NS m
†Quemisia gravis NS
a Museum and collector abbreviations are as follows: AMNH, American Museum of Natural History; CML, Universidad Nacional de Tucumán, Colección de Mamíferos Lillo; FMNH, Field
Museum of Natural History; MCNM, Museu de Ciências Naturais, Mammals Collection, Pontifícia Universidade Católica Minas Gerais; MCNU, Museu de Ciências Naturais da Universidade
Luterana do Brasil; MEPN, Museo Escuela Politécnica Nacional, Ecuador; MVZ, University of California, Berkeley, Museum of Vertebrate Zoology; MN, Universidade Federal do Rio de Janeiro,
Museu Nacional; ROM, Royal Ontario Museum; UFMG, Universidade Federal de Minas Gerais, Mammals Collection; UFROM, Universidade Federal de Rondônia, Mammals Collection; USNM,
United States National Museum of Natural History; ZTNH, University of Vermont, Zadock Thompson Natural History Collections; LGA, collector number of Ana Paula Carmignotto and Roberta
Paresque; LHE, collector number of Louise H. Emmons; RBP, Rafael Borroto-Paez collection (and specimen number); RM, collector number Raquel Moura.
b Chimera of Pedetes species capensis (c) and surdaster (s)
c Chimera of Ctenodactylus species gundi (g) and vali (v)
d Mislabeled 12S rRNA and GHR sequences on GenBank: as Coendou bicolor, but these represent C. prehensilis (R. Voss, in litt., 6 Mar 2012)
e Our attempts failed to amplify DNA from a ~500-year old bone specimen of Cuscomys oblivatus (YPM 3318; Machu Picchu). There were reports in 2009 of a Cuscomys individual sighted at
Machu Picchu (L. Emmons in litt , 1 Aug 2011)
f Species was described by de Frietas et al. (2012)
g Echimys saturnus was recorded at Tiputini Biodiversity Reserve in 2005 (Blake et al., 2010)
h Mislabeled cyt-b and 12S rRNA sequences on GenBank: as Isothrix bistriata but these are I. negrensis (Patterson and Velazco, 2008)
i Mislabeled cyt-b sequence on GenBank: as Makalata didel phoides, but this is M. macrura (Patterson and Velazco, 2008)
j The name Makalata obscura cannot be assigned to a speciﬁc population; holotype specimen is lost with an unknown locality (Emmons, 2005)
k The published cyt-b sequence of Phyllomys unicolor (EF608188) is a mitochondrial numt of P. pattoni (C. Loss in litt, 30 Nov 2012)
l Amblyrhiza is allied outside of Chinchilloidea based on basicranial characters (MacPhee, 2011)
m Elasmodontomys is allied outside the Echimyidae + Capromyidae clade (including Myocastor) based on basicranial characters (MacPhee, 2011)
Caviomorph evolution through time
Appendix 2. Detailed justication of the 22 fossil calibration points used in this study. Three additional calibration points
were rejected because of the temporal inconsistencies they introduced during cross-validation analyses (see Rejected Cali-
brations below). Each entry begins with the constrained clade and its corresponding code (A-Y; see Fig. 3), then the lognor-
mal prior’s upper 95% range in millions of years (Ma) and its shape (mean, SD) as implemented in BEAST (an “R”next to the
mean refers to the mean in “real space”). Each prior was oset to the minimum age of the oldest crown fossil assigned to that
clade, and a soft maximum age set at the lower 5% of the prior. We then list fossil taxa supporting the minimum age esti-
mates, museum voucher specimen (if any), role in our dating scheme, and references for their ancestry; we also include the
geological formation used in minimum age estimates, aging method, and associated references. Horizons used in maximum
age estimates and their references conclude each entry. SALMA refers to South American Land Mammal Age; see referenced
publications for museum abbreviations and other details.
A - Ctenohystrica: 46.0–65.8 Ma (1.341,1)
Based on †Chapattimys wilsoni, the earliest condently placed stem ctenodactyloid: 46–51 Ma, early-to-
middle Eocene (Marivaux et al., 2004; Sallam et al., 2009; Sallam et al., 2011; Antoine et al., 2012). From
the Subthu group, Kuldana Formation, India and Pakistan (Flynn et al., 1986; Antoine et al., 2012).
Maximum: to the occurrence of eurymylids, e.g., †Heomys, a primitive rodent from the early Paleocene,
65.8 Ma (Hussain et al., 1978; Meredith et al., 2011).
B - Diatomyidae–Ctenodactylidae: 28.3–46.0 Ma (1.230,1)
Based on †Fallomus razae, †Fallomus ginsburgi, and †Fallomus quraishyi (GSP 21218, DC 683, DC 438),
the oldest stem diatomyid; crown member of Ctenodactylidae-Diatomyidae: >28.3 Ma, Rupelian stage,
early Oligocene (Dawson et al., 2006). From the Bugti Hills, Paali nala C2, Chitarwata Formation,
Balochistan, Pakistan (Flynn et al., 1986; Marivaux and Welcomme, 2003). Maximum: to the occurrence
of †Chapattimys, 46 Ma (Flynn et al., 1986; Antoine et al., 2012).
C - Hystricognathi: 41.0–56.0 Ma (1.063,1)
Based on †Cachiyacuy contamanensis, †Cachiyacuy kummeli, †Canaanimys maquiensis, †Gaudeamus hylaeus,
and †Gaudeamus aslius (MUSM 1871, MUSM 1882, MUSM 1890, CGM 66007, CGM 66006), the
oldest crown hystricognaths in South America (see calibration E), Barrancan SALMA, middle Eocene. e
oldest crown members in Africa (Gaudeamus) are younger at ~34 Ma (Sallam et al., 2009; Sallam et al.,
2011; Antoine et al., 2012). From the CTA-27 Locality, Yahuarango Formation, Contamana, Loreto, Peru;
Jebel Qatrani Formation, Fayum Depression, Egypt (Sallam et al., 2009; Sallam et al., 2011; Antoine et al.,
2012). Maximum: to the early Eocene (56 Ma), the earliest proposed date for the Santa Rosa fauna, Peru
(Frailey and Campbell, 2004; Meredith et al., 2011).
D - Hystricidae: 11.0–15.0 Ma (R1.280,1)
Based on Atherurus indet (SA 64), the oldest condently dated crown Hystricidae fossil: –11 Ma, early
Vallesian (=MN 9), late Miocene(Mein and Pickford, 2006). From the Sheikh Abdallah, Western Desert,
Egypt (Mein and Pickford, 2006). Maximum: to base of the Astaracian– 15 Ma (=MN 6), because of the
putative Atherurus fossil (†A. karnuliensis, formerly †Sivacanthion complicatus) from the Lower Silwaliks of
Pakistan (Colbert, 1935; van Weers, 2005).
E - Caviomorpha – Phiomorpha: 41.0–56.0 Ma (1.063,1)
Based on †Cachiyacuy contamanensis; †Cachiyacuy kummeli; †Canaanimys maquiensis (MUSM 1871, MUSM
1882, MUSM 1890), the oldest known caviomorphs in South America; stem Caviomorpha, not referable
to any superfamily: ~41 Ma (40.94–45.94 Ma radiometric; 40.94–41.6 Ma including bio-chronology),
Barrancan SALMA, middle Eocene (Antoine et al., 2012). From the CTA-27 Locality, Yahuarango
Formation, Contamana, Loreto, Peru. Radiometric age (40Ar/39Ar) and mammalian biochronology (Antoine
et al., 2012). Maximum: to early Eocene (56 Ma), in accordance with the earliest proposed date for the
Santa Rosa fauna, Peru (Frailey and Campbell, 2004; Meredith et al., 2011).
F - Cavioidea – Erethizontoidea: 31.3–41.0 Ma (0.63,1)
Based on †Andemys termasi (SGOPV 2933), oldest stem member of Cavioidea sensu stricto; may instead be
basal members of Dasyproctidae: 31.3–33.6 Ma, Tinguirirican SALMA, late Eocene-early Oligocene (Wyss
et al., 1993; Bertrand et al., 2012; Pérez and Pol, 2012;). From the Abanico (= Coya-Machalí) Formation,
Upham and Patterson
Tinguiririca River valley, Termas del Flaco, central Chile. Radiometric age: 40K/40Ar (Wyss et al., 1990;
Flynn et al., 2003; Dunn et al., 2013). Maximum: to the earliest caviomorph fossils in South America, ~41
Ma (Antoine et al., 2012).
G - Cavioidea (=Caviidae, Cuniculidae, Dasyproctidae): 24.2–30.77 Ma (0.237,1)
Based on †Asteromys punctus and †Chubutomys simpsoni, oldest crown members of Cavioidea sensu stricto
(“Eocardiidae”; stem members of Caviidae): 24.2–29.4 Ma, Deseadan SALMA, late Oligocene (Pérez and
Pol, 2012). From Cabeza Blanca and Lagunade los Machos, Sarmiento Formation, Argentina (Pérez and
Vucetich, 2011; Dunn et al., 2013). Maximum: to the base of the youngest assemblage that lacks crown
cavioid fossils, which is La Cantera, Gran Barranca, Argentina, 30.77 Ma (Vucetich et al., 2010; Pérez and
Pol, 2012; Dunn et al., 2013).
H - Caviidae (= Hydrochoerus, Kerodon, Dolichotis, Galea, Microcavia, Cavia): 11.8–13.8 Ma (0.640,1)
Based on†Prodolichotis pridiana, the oldest crown member of Caviidae: 11.8–13.5 Ma, Laventan SALMA,
middle Miocene (Pérez and Pol, 2012). From the La Victoria and Villa Vieja formations, La Venta section of
Colombia (Fields, 1957; Kay et al., 1999). Maximum: to the start of the Colloncuran SALMA (13.8 Ma),
since cavioid fossils of that period (e.g., †Guiomys unica) are outside of crown Caviidae (Pérez and Pol, 2012).
I - Hydrochoerus – Kerodon: 6.1–11.8 Ma (0.095,1)
Based on †Cardiomys cavinus and †Cardiatherium chasicoense, the oldest crown hydrochoerines: 6.1–9.07
Ma, Chasicoan SALMA, late Miocene (Pérez and Pol, 2012). From the Arroyo Chasicó Formation, central
Argentina. Radiometric age: correlated with Loma de Las Tapias Formation in northwestern Argentina
(Deschamps et al., 2009; Pérez and Pol, 2012). Maximum: to the start of the Laventan SALMA (11.8 Ma)
since the cavioids of this period (e.g., †Prodolichotis) are outside of crown Hydrochoerinae. is is the youngest,
well-sampled rodent assemblage lacking forms potentially allied to Hydrochoerinae (Pérez and Pol, 2012).
J - Microcavia / Cavia – Galea: 6.1–11.8 Ma (0.095,1)
Based on †Allocavia chasicoense, oldest crown caviine: 6.1–9.07 Ma, Chasicoan SALMA, late Miocene
(Pérez and Pol, 2012). From the Arroyo Chasicó Formation, central Argentina. Radiometric age: correlated
with Loma de Las Tapias Formation in northwestern Argentina (Pascual, 1962). Maximum: to the start of
the Laventan SALMA (11.8 Ma) since the cavioids of this period (e.g., †Prodolichotis) are outside of crown
Caviinae (Pérez and Pol, 2012).
K - Microcavia – Cavia: 4.0–9.07 Ma (R1.614,1)
Based on †Paleocavia impar, the oldest stem taxon to Microcavia: 4.0–5.3 Ma, Montehermosan SALMA, early
Pliocene (Pérez and Pol, 2012). From the Monte Hermoso Formation, Argentina (Ameghino, 1889; Cione
and Tonni, 1995; Schultz et al., 2002). Maximum: to the base of the upper section of the Arroyo Chasicó
Formation (9.07 Ma) to take into account possibly older fossils allied to this group in the Huayquerian (Pérez
and Pol, 2012).
L - Chinchillidae: 9.07–19.04 Ma (0.655,1)
Based on †Prolagostomus sp. (AMNH DVP 99300; 9587 †P. imperialis), oldest crown members of
Chinchillidae; stem members of the Lagostominae: 9.07–19.04 Ma, Pinturan SALMA, early-to-late Miocene
(Kramarz, 2002; Croft et al., 2011). From (youngest occurrence) Arroyo Chasicó Formation, central Argentina
and (oldest occurrence) Pinturas Formation, Argentina (Bondesio et al., 1980; Kramarz, 2002; Dunn et al.,
2013). Maximum: to the base of the Pinturan SALMA (19.04 Ma) based on the earliest occurrence of
†Proglagostomus in the Pinturas Formation of southern Argentina (Kramarz, 2002; Dunn et al., 2013).
M - Chinchilloidea – Octodontoidea: 31.3–41 Ma (0.627,1)
Based on †Eoviscaccia frassinettii (SGOPV 2935), oldest stem member of Chinchilloidea: 31.3–33.6 Ma,
Tinguirirican SALMA, late Eocene-early Oligocene. Eoviscaccia is older than the oldest stem octodontoid,
†Draconomys verai, from 30.62–30.77 Ma at La Cantera, Argentina (Wyss et al., 1993; Vucetich et al.,
2010; Bertrand et al., 2012). From Abanico (= Coya-Machalí) Formation, Tinguiririca River valley, Termas
del Flaco, central Chile. Radiometric age: 40K/40Ar (Wyss et al., 1990; Flynn et al., 2003; Dunn et al., 2013).
Maximum: to the earliest caviomorph fossils in South America ~41 Ma (Antoine et al., 2012).
Caviomorph evolution through time
N - Abrocomidae: 2.0–6.1 Ma (R1.305,1)
Based on Abrocoma (MMP 1059-M, MACN19722), oldest crown members of Abrocomidae (Abrocoma +
Cuscomys): >2.0 Ma, Sanandresian substage, Upper Marplatan SALMA, late Pliocene (Verzi and Quintana,
2005b; Verzi et al., 2013). From Punta San Andrés and Santa Isabel, Buenos Aires, Argentina. Biochrono-
logical and magnetostratigraphic age (Verzi and Quintana, 2005b). Maximum: to the start of the Huay-
querian SALMA (6.1 Ma) based on the rst occurrence of the stem abrocomid, †Abrocoma (Protoabrocoma)
antiqua, in the late Miocene of Bolivia and western Argentina (Cione et al., 2000; Verzi and Quintana,
2005b; Verzi et al., 2013).
P - Octodontidae: 6.8–11.8 Ma (R1.590,1)
Based on †Pseudoplateomys innominatus (MACN 8363), the oldest crown octodontid (or octodontine sensu
Verzi et al., 2013): >6.8 Ma, Huayquerian SALMA, late Miocene (Verzi et al., 2013). From Quebrada de
La Troya, La Rioja, Argentina. Radiometric age (Ciccioli, Limarino and Marenssi, 2005). Maximum: to the
start of the Laventan SALMA (11.8 Ma), beyond which no crown octodontid fossils are found, only stem
fossils such as †Acarechimys (Verzi et al., 2013).
Q - Aconaemys / Spalacopus – Octodon: 5.0–11.8 Ma (0.272,1)
Based on †Pithanotomys (MACN-A 1648, †P. columnaris holotype), crown octodontid, sister to Aconaemys:
>5.0 Ma, Montehermosan SALMA, early Pliocene (Verzi et al., 2013). From Farola Monte Hermoso,
Buenos Aires, Argentina. Biochronological age (Zárate et al., 2005; Verzi, 2008). Maximum: to the start of
the Laventan SALMA (11.8 Ma), beyond which no crown octodontid fossils are found, only stem fossils
such as †Acarechimys (Verzi et al., 2013).
R - Octomys – Pipanacoctomys / Tympanoctomys / Salinoctomys: 2.0–6.8 Ma (R1.528,1)
Based on †Abalosia castellanosi (PVL 1252, †Plataeomys castellanosi holotype), oldest member of the desert-
adapted clade in Octodontidae: >2.0 Ma, Sanadresian substage, Upper Marplatan SALMA, late Pliocene
(Verzi et al., 2013). From Punta San Andrés and Santa Isabel, Buenos Aires, Argentina. Biochronological
and magnetostratigraphic age (Verzi and Quintana, 2005a). Maximum: to the minimum age for crown
Octodontidae (6.8 Ma), based on †Pseudoplateomys (Verzi et al., 2013).
S - Ctenomys: 3.5–5.3 Ma (R0.573,1)
Based on †Ctenomys uquiensis (MLP 96-II-29-1), the oldest member of Ctenomys crown group: ~3.5
Ma, Chapadmalalan SALMA, late Pliocene (Verzi et al., 2013). From Esquina Blanca, Jujuy, Argentina.
Radiometric andmagnetostratigraphic age (Reguero et al., 2007; Verzi et al., 2010). Maximum: to the last
occurrence of the Ctenomys sister taxon, †Praectenomys rhombidens, 5.3 Ma, from the Umala Formation,
Bolivia (Quintana, 1994; Verzi and Montalvo, 2008; Verzi et al., 2013).
V - Echimyidae “arboreal clade”: 15.7–24.2 Ma (0.495,1)
Based on †Maruchito trilofodonte (MLP 91-IV-1-22), crown echimyid; oldest stem taxon to Echimys + Phyllomys:
~15.7 Ma, Colloncuran SALMA, middle Miocene (Verzi et al., 2013). From Cañadón del Tordillo, Neuquén,
Argentina. Radiometric age (Vucetich et al., 1993; Madden et al., 1997). Maximum: to the start of the
Deseadan SALMA, 24.2 Ma (Dunn et al., 2013).
W - Thrichomys – Myocastor / Callistomys / Hoplomys / Proechimys: 6.0–11.8 Ma (0.113,1)
Based on †Pampamys (GHUNL-Pam 2214), crown echimyid; oldest stem taxon to richomys: 6.0–9.3 Ma,
Chasicoan - Huayquerian SALMA, late Miocene (Olivares et al., 2012; Verzi et al., 2013). From Laguna Chillué,
Cerro Azul Formation, Bajo Giuliani, Telén, Loventué, La Pampa, Argentina. Biochronological age (Zárate et
al., 2005; Verzi et al., 2008). Maximum: to the start of the Laventan SALMA, 11.8 Ma (Kay et al., 1997).
X - Myocastor – Callistomys / Hoplomys / Proechimys: 6.0–11.8 Ma (0.113,1)
Based on Myocastor (MACN 5404, †M. columnaris holotype), crown echimyid; oldest Myocastor species:
>6.0 Ma, Huayquerian SALMA, late Miocene (Verzi et al., 2013). From the “Mesopotamiense” or “Con-
glomerado osífero”, Ituzaingó Formation, Entre Ríos, Argentina. Biochronological age (Cione et al., 2000;
Candela and Noriega, 2004; Zárate et al., 2005). Maximum: to the start of the Laventan SALMA, 11.8 Ma
(Kay et al., 1997).
Upham and Patterson
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U - Capromyidae: 14.86–29.4 Ma (1.032,1)
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as stem isolobodontine from anity of three teeth with †Isolobodon from Hispaniola: >14.86 Ma, early
Miocene (MacPhee et al., 2003; MacPhee, 2005). From the Lagunitas Fm (Cuba), Domo de Zaza, Sancti
Spiritus. Radiometric age: Rb-Sr (also older results that compare to Santacrucian SALMA) (MacPhee,
2005). Maximum: to the base of the Deseadan SALMA, 29.4 Ma (Dunn et al., 2013).
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Appendix 3. Comparison of substitution saturation across codon positions in 1140 bp of the cytb mtDNA gene. Plotted are
pairwise comparisons among taxa for 1. numbers of transitions (Ti) and transversions (Tv) and their corresponding linear
models, 2. number of transitions versus percent sequence divergence (uncorrected–p) for all cyt–b codon positions, and 3)
number of transversions versus cyt–b percent sequence divergence.