Updated and annotated checklist of recent mammals from Brazil

Abstract

An updated and annotated checklist of mammals occurring in Brazil is presented. A total of 751 native species, distributed in 249 genera, 51 families and 11 orders were recorded to the country. The Brazilian mammalian fauna shows an elevated rate of endemism (30%; 223 species). Among the species evaluated by IUCN (668 species; 90%), a total of 80 (10.6% of total mammalian fauna) are Threatened, 28 (3.9%) are considered as Near Threatened, two species (0.3%) are presumable Extinct, 96 (12.8%) are considered with Deficient Data for conservation and 462 (61.6%) are considered as Least Concern. Fifteen new species were described since the last national compilation (published in 2017), which associated to new records to the country and synonimizations resulted in an increment of 30 species. Eight non-native species were introduced to the country, including the recently established Asiatic cervids Rusa unicolor (sambar) and Axis axis (chital). Seven native species (five primates and two hystricomorph rodents) have been translocated from their areas of natural occurrence to other areas inside the country.

Key words
biodiversity; conservation; Mammalia; Neotropical region; taxonomic list

INTRODUCTION

Currently, more than 6,400 mammal species are recognized worldwide. The Neotropics is the most diverse biogeographic realm, comprising around 25% of the mammalian diversity (Burgin et al. 2018). Within the Neotropical realm, Brazil is the largest country, covering an area nearly half of South America and housing two of the hotspots for the world conservation of biodiversity (Cerrado and Atlantic Forest) (Myers et al. 2000), as well as a great part of the Amazon Forest. Some authors (Mittermeier et al. 1997, Costa et al. 2005) have considered Brazil as the country with the highest mammalian diversity in the world.

Considerable attempts have been made to dimension the Brazilian mammalian diversity (Fonseca et al. 1996, Vivo 1996, Costa et al. 2005, Reis et al. 2006, 2011, Paglia et al. 2012, Percequillo & Gregorin 2017) and each new list adds new species as the result of a crescent effort on field sampling and taxonomic revisions (Fonseca et al. 1996, Paglia et al. 2012). The last compilation of mammals from Brazil (Percequillo & Gregorin 2017) listed a total of 721 species (or 778 valid nominal taxa, including subspecies). This list was produced as part of a joint effort of the Brazilian Society of Zoology aiming to produce an online database of the Brazilian fauna. It brought an accretion of 20 species in relation to the previous list of mammals occurring in Brazil (Paglia et al. 2012) and represents a very useful reference for the diversity of the group in the country. However, several new taxa described posteriorly to Paglia et al. (2012) were omitted by Percequillo & Gregorin (2017) (e.g. Hrbek et al. 2014, Feijó et al. 2015, Moratelli & Dias 2015, Christoff et al. 2016, Pavan et al. 2017). Moreover, some taxonomic revisions resulting in nomenclatural changes conducted posteriorly to Paglia et al. (2012) annotated list (e.g. Berta & Churchill 2012, Feijó & Cordeiro-Estrela 2016, Nascimento & Feijó 2017) were not considered by Percequillo & Gregorin (2017). Still, progress in alpha-taxonomy has already been made after the compilation of Percequillo & Gregorin (2017) (e.g. Percequillo et al. 2017, Boubli et al. 2018, Garbino et al. 2019, Miranda et al. 2018, Feijó et al. 2018, Serrano-Villavicencio et al. 2019, Pavan 2019, Feijó et al. 2019). These three factors drew attention to the need for a reassessment of the mammalian diversity occurring in Brazilian territory.

Information on alpha-taxonomy is crucial in this current period of accelerated loss of biodiversity (Grieneisen et al. 2014). In this view, lists of species are of great importance for knowledge on regional diversity and the development of conservation plans. Herein, we compile all the information available about the valid mammalian species occurring in Brazil to the present date, aiming to produce an updated and annotated list for the country. This new compilation presents all new taxa described since the publication of Paglia et al. (2012) and Percequillo & Gregorin’s (2017) lists as well as taxonomic modifications occurred since then, accompanied by justifications based on recent findings. We also included available and unpublished data on the status of exotic and invasive mammalian species in Brazil.

MATERIALS AND METHODS

The present annotated checklist was produced based on the two previous published lists of Brazilian mammals (Paglia et al. 2012, Percequillo & Gregorin 2017). An extensive research on all information (scientific articles, books and book chapters) published from 2012 to August 2019 was made on platforms Google, Google Scholar, Scopus and Web of Science, using as search terms the name of each genus and family presented on each previous lists (Paglia et al. 2012, Percequillo & Gregorin 2017) associated to “Brazil” and “New” (examples: “Didelphidae Brazil”, “new Didelphidae”, “Monodelphis Brazil”, “new Monodelphis”). The search terms “Sigmodontinae Brazil”, “new Sigmodontinae”, “new genus mammal Brazil” and “new genus Mammalia Brazil” were also applied. The contents of each genus were also compared with the Mammal Diversity Database (www.mammaldiversity.org) (ASM 2019). We also checked the Brazilian institutional platform of academic productivity (Plataforma Lattes, accessed on: lattes.cnpq.br) for the latest scientific articles published by the main mammalian taxonomists in activity in Brazil. All the new taxa described after the publication of the latest list (Percequillo & Gregorin 2017) were added to this new list together with information on type locality and distribution. The taxonomic modifications as well as new evidences for taxa revalidation or refutation occurred after the publication of Percequillo & Gregorin’s (2017) list were included as ‘Remarks’ containing justifications based on the respective researches. We have also included as ‘Remarks’ observations on forms with unstable taxonomy investigated in the last two years, even when in nomenclatural concordance with Percequillo & Gregorin (2017). Information on status of conservation of all species was given based on the latest evaluation of the International Union for the Conservation of Nature (IUCN) and on the Brazilian Red List (ICMBio 2018). The species endemic to the country were also indicated.

Finally, we present a section including the list of all allochthonous species so far recorded living in the wild in natural environments in Brazil, as well as the native displaced species. That information was obtained by published data (articles, books, and book chapters), unpublished data (doctoral thesis, master’s dissertations, meeting communications) and personal communications of researchers. We included only wild non-native species, excluding feral and free-roaming domestic species (Canis familiaris, Felis catus, Bos sp., Equus sp., Capra sp.). To know more about these domestic species, see Rosa et al. (2017a).

RESULTS

A total of 751 autochthonous mammalian species, distributed in 249 genera, 51 families and 11 orders were assigned to Brazil. The most speciose order is Rodentia with 258 species, followed by Chiroptera (182 spp.), Primates (126 spp.), Didelphimorphia (62 spp.), Artiodactyla (57 spp.), Carnivora (36 spp.), Cingulata (12 spp.), Pilosa (12 spp.), Lagomorpha (2 spp.), Perissodactyla (2 spp.), and Sirenia (2 spp.). A total of 80 species (10.6%) are included in some global threat category (12 Critically Endangered, 31 Endangered and 37 Vulnerable), 28 (3.7%) are considered as Near Threatened, two species (0.3%) are presumable Extinct, 96 (12.8%) are considered with Deficient Data for conservation and 462 (61.6%) are including in Least Concern (non-threatened) category (IUCN 2019). Eighty-three species (11%) are pending (re)evaluation due to recent taxonomic rearrangements or new species descriptions. According to the Brazilian red list (ICMBio 2018), 110 taxa (15%) are listed under some threat category (10 species are Critically Endangered, 41 Endangered, and 52 Vulnerable). A total of 223 species (30%) are endemic to Brazil (Appendix).

REMARKS

Order Didelphimorphia

• 1. Gracilinanus peruanus (Tate, 1931) was resurrected from the synonym of G. agilis (originally Marmosa gracilis peruana Tate, 1931) based on phylogenetic analyses of cytochrome b (Cytb) sequences and external and craniodental morphology (Semedo et al. 2015). The occurrence of this taxon in Brazil was assigned to the states of Rondônia and Mato Grosso (Semedo et al. 2015).

• 2. Marmosa (Micoureus) phaea Thomas, 1899 was recorded for the right bank of Rio Tiquié, Comunidade Colina, Amazonas state (0.12° N, 69.01° W) (Voss et al. 2020).

• 3. Voss et al. (2020) based on molecular and morphological evidence resurrected Marmosa (Micoureus) rapposa Thomas, 1899 (type locality: Huadquiña, Cuzco, Peru). This species was previously treated a junior synonym or subspecies of Marmosa (Micoureus) regina Thomas, 1898. In Brazil, M. rapposa is recorded in Mato Grosso and Mato Grosso do Sul states (Voss et al. 2020).

• 4. Marmosa (Micoureus) rutteri Thomas, 1924 (type locality: “Tushemo, near Masisea”, Ucayali, Peru) is recognized as a valid species by Voss et al. (2020) apart from Marmosa (Micoureus) regina Thomas, 1898. Marmosa (M.) rutteri is distributed in the lowland Amazon forest of southeastern Colombia, eastern Ecuador, eastern Peru and western Brazil. Records in Brazil include Acre and Amazonas states.

• 5. In the subgeneric classification of Marmosa of Voss et al. (2014a), Marmosa lepida Thomas, 1888 is placed within Stegomarmosa Pine, 1972.

• 6. Díaz-Nieto et al. (2016) performed phylogenetic analysis on Cytb sequences of all recognized Marmosops species and found an internal dichotomy, leading to a subgeneric classification.

• 7. Marmosops impavidus (Tschudi, 1845) was considered a nomen dubium by Díaz-Nieto et al. (2016: 931) and therefore it was not included herein.

• 8. Voss et al. (2019) recognized Metachirus myosuros (Temminck, 1824) (type locality: Ipanema, São Paulo, Brazil) as a species apart from M. nudicaudatus based on cranial traits and phylogenetic analyses of the Cytb. Metachirus nudicaudatus is now restricted to the Guiana Shield region with confirmed records from Amapá state in Brazil. Voss et al. (2019) reported specimens of M. myosuros from the Brazilian states of Acre, Amazonas, Bahia, Espírito Santo, Minas Gerais, Rio de Janeiro, São Paulo, and Rondônia.

• 9. Pavan et al. (2016), analyzing sequences of Cytb and four nuclear genes, recovered monophyletic species groups within Monodelphis. Subsequently, Pavan & Voss (2016) formally organized such groups into subgenera, showing diagnostic morphological characters.

• 10. Monodelphis (Microdelphys) gardneri Solari, Pacheco, Vivar & Emmons, 2012 was described from specimens collected in the montane forests of central Peru and there is no confirmed record for Brazil. The species is not included in the present list.

• 11. Monodelphis (Monodelphis) vossi was described from three specimens collected in two savanna localities in Roraima state (Pavan 2019). Referred as Monodelphis “species 3” in Pavan et al. (2016) and Pavan & Voss (2016).

• 12. Monodelphis (Mygalodelphys) handleyi Solari, 2007 was reported from Brazil based on one individual collected in an Amazonian savanna area, Humaitá, Amazonas (Bezerra et al. 2019).

• 13. Monodelphis (Mygalodelphys) saci was described from 18 specimens collected in Brazilian Amazon (states of Pará, Mato Grosso, Rondônia and Acre) (Pavan et al. 2017). Referred as Monodelphis “species 2” in Pavan et al. (2016) and Pavan & Voss (2016).

• 14. Voss et al. (2018) revised Philander through mitochondrial and nuclear genes phylogeny and morphological analyses and considered eight valid species. According to these authors, Philander canus (Osgood, 1913) (type locality: Moyobamba, San Martín, Peru) is the name applicable to the form occurring in western Brazil (Cerrado biome).

• 15. According to Voss et al. (2018), Philander quica (Temminck, 1824) is the name applicable to the Brazilian Atlantic Forest form, considering that the holotype of P. frenatus (Olfers, 1818) was collected in eastern Amazonia. Voss et al. (2018) considered P. frenatus (Olfers, 1818) a junior synonym of P. opossum (Linnaeus, 1758).

• 16. Philander pebas was described from specimens collected on the left bank of the Rio Juruá, state of Amazonas. In Brazil, it is referred to Amazon Forest habitats of Amazonas and Acre states (Voss et al. 2018).

Order Sirenia

• 1. Trichechus pygmaeus Van Roosmalen et Van der Vlist, 2015 was described based on the holotype (CCM181) and an alive individual kept in semi-captivity. According to Van Roosmalen (2015), T. pygmaeus differs from T. inunguis by overall smaller size, darker coloration, skull morphology, and fewer cheek teeth. In addition, Van Roosmalen (2015) obtained a fragment of 410 bp of the left domain of the mitochondrial control region (D-loop) from the living specimen and recovered a haplotype found by Garcia-Rodriguez et al. (1998) for T. inunguis. Still, Van Roosmalen (2015) interpreted this result as a possible consequence of the slow mutation rate of the control region in manatees, or even a past event of isolation of T. pygmaeus followed by the hybridization with T. inunguis. The specific status of T. pygmaeus was contested by authorities who claimed that specimens and individuals attributed to this taxon are actually juvenile T. inunguis (see Jefferson et al. 2015). Therefore, herein we did not consider T. pygmaeus as valid species.

Order Cingulata

• 1. Feijó et al. (2019) performed phylogenetic analyses using two mitochondrial markers (Cytb and cytochrome oxidase subunit I [COI]) and one nuclear marker (exon 28 of the von Willebrand factor [vWF]) and recovered three clades within Dasypus, which were treated as subgenera (Dasypus, Hyperoambon and Muletia).

• 2. Feijó & Cordeiro-Estrela (2016) revised Dasypus kappleri using morphological and morphometric (linear and 2D geometric) analyses and considered it a species complex, which was further supported by phylogenetic analyses (Feijó et al. 2019). Dasypus kappleri beniensis Lönnberg, 1942 was elevated to full species. In Brazil, this taxon is known from the states of Pará, Rondônia, and Mato Grosso (Amazon biome).

• 3. Feijó & Cordeiro-Estrela (2016) restricted Dasypus kappleri Krauss, 1862 to the form occurring to the north of Amazon River (Guiana shield) in French Guiana, Suriname, Guyana, eastern Venezuela and Brazilian states of Amapá, Amazonas and Pará.

• 4. Feijó & Cordeiro-Estrela (2016) resurrected Dasypus pastase (Thomas, 1901) (type locality: “Sarayacu, upper Pastasa River”, Pastaza, Ecuador) to the form occurring on the foothills of the eastern Andes in Peru, Ecuador, Colombia, Venezuela (south of the Orinoco River) and Brazilian state of Amazonas (south of Javari river).

• 5. Feijó et al. (2018) revised the genus Dasypus through qualitative morphological analysis and linear and 2D geometric morphometrics, recognizing Dasypus hybridus (Desmarest, 1804) as a subspecies of Dasypus septemcinctus Linnaeus, 1758. Molecular phylogeny and species delimitation analyses also supported this arrangement (Feijó et al. 2019).

• 6. Gibb et al. (2015) erected family Chlamyphoridae Bonaparte, 1850 to include subfamilies Euphractinae, Chlamyphorinae and Tolypeutinae, a clade highly divergent from Dasypus clade in the mitogenomic phylogenetic analysis performed by the authors. Dasypodidae was restricted to Dasypus genus.

• 7. The occurrence of Cabassous chacoensis Wetzel, 1980 in Brazil is uncertain. In its description, Wetzel (1980) referred to one alleged specimen from Brazil obtained from the Buenos Aires Zoo in 1904. Besides that, no other individual of this species was ever recorded in the country. Therefore, considering there is no reliable evidence of this species in Brazil and following recent reviews (Wetzel et al. 2008, Hayssen 2014, Brandão et al. 2019), we did not consider C. chacoensis as part of the Brazilian mammalian fauna.

• 8. Feijó & Langguth (2013) treated Cabassous squamicaudis (Lund, 1845) (type locality: “Rio das Velhas Floddal, Lagoa Santa, Minas Gerais, Brazil”) as full species apart from Cabassous unicinctus (Linnaeus, 1758) based on phenotypic differences, which has been corroborated by molecular studies (Schetino M.A.A., unpublished data).

Order Pilosa

• 1. Miranda et al. (2018) revised Cyclopes through morphological, morphometric and molecular data (Cytb, COI and mitochondrial control region [CR]) and recognized seven valid species.

• 2. Miranda et al. (2018) designated a neotype for C. didactylus (Linnaeus, 1758) (FMNH 93175, type locality: Kayser-Gebergte Airstrip, Suriname) and cited the occurrence of this taxon to Suriname, Venezuela, French Guiana, Brazil and Trinidad and Tobago. In Brazil, C. didactylus occurs disjointly in Amazon (states of Amazonas, Pará, Maranhão and Piauí) and northeast Atlantic Forest (states of Rio Grande do Norte, Paraíba, Pernambuco and Alagoas).

• 3. Cyclopes ida Thomas, 1900 was originally described as a subspecies of C. didactylus. Miranda et al. (2018) elevated ida to species level and assigned its distribution to Ecuador, Colombia, Peru and Brazilian states of Acre and Amazonas.

• 4. Cyclopes rufus (type locality: “Porto Velho, Rondônia, Brazil”) was described by Miranda et al. (2018) from specimens collected in the Brazilian state of Rondônia, between Madeira and Aripuanã rivers.

• 5. Cyclopes thomasi (type locality: “Porto Walter, Acre, Brazil”) was described by Miranda et al. (2018) based on specimens from few localities in Peru and Brazilian states of Acre and Amazonas.

• 6. Cyclopes xinguensis (type locality: Vitória do Xingu, Pará, Brazil [Usina Belo Monte]) was described by Miranda et al. (2018) from specimens collected in the Brazilian states of Pará and Amazonas.

Order Primates

• 1. The status of Cebuella niveiventris Lönnberg, 1940 as a full species, herein adopted, is resultant from phylogenetic analysis of of Cytb sequences and reduced-representation genome sequencing (ddRADseq) (Boubli et al. 2018), which was further supported by molecular species delimitation analysis (Garbino et al. 2019).

• 2. Mico munduruku Costa-Araújo, Farias, & Hrbek, 2019 (type locality: Boca do Crepori community (0546’55’S, 5715’14’W), right margin of the mouth of the Crepori River, Itaituba municipality, Pará State, Brazil) was described from specimens collected in seven localities in the Amazon forest, Pará state, between Jamanxim, Novo, Tapajós and Cururú rivers (Costa-Araújo et al. 2019).

• 3. The capuchin monkeys are traditionally classified in two groups: gracile (or untufted) and robust (or tufted) species. Silva-Júnior (2002) treated these two groups as subgenera (Cebus for gracile and Sapajus for robust) of Cebus. Later, Lynch Alfaro et al. (2012a, b, 2014) advocate for the use of Cebus and Sapajus as distinct genera, based on morphological, genetic, biogeographic, behavioral, and ecological differences. This new classification has provoked a heated debate (Rosenberger 2012, Feijó & Langguth 2013, Garbino 2015, Gutiérrez & Marinho-Filho 2017). Because there is no objective and universal criteria for defining genera, Garbino (2015) proposed the use of an integrative approach based on multiple sources of evidence to define genera. Herein, we considered Cebus Erxeleben, 1777 and Sapajus Kerr, 1792 as valid genera.

• 4. Boubli et al. (2008) described Cacajao hosomi (type locality: Imeri Mountains, Serra do Xamatá, Amazonas, Brazil) and C. ayresi (type locality: right bank of lower Rio Aracáy, Amazonas, Brazil) apart from C. melanocephalus (Humboldt, 1811) based on pelage coloration and divergences of Cytb sequences. Later, Ferrari et al. (2014) considered C.ayresi as a subspecies of C. melanocephalus and resurrected C. ouakary as a valid species. Ferrari et al. (2014) also refer the nominal subspecies C. m. melanocephalus as Neblina black-faced uacari, which is the designation given by Boubli et al. (2008) to C. hosomi. Surprisingly, Ferrari et al. (2014) did not mention hosomi along the full main text but it is implicit that C. m. melanocephalus, as established there, includes hosomi. Ferrari et al. (2014) also argued that pelage differentiation between ayresi and melanocephalus (including hosomi) represents a natural geographic gradient from darker to lighter patterns while the genetic distances between the two clades is far below than interspecific distances observed in other mammalian lineages, including other primate genera. Moreover, Ferrari et al. (2014) call attention to the utilization of a single mitochondrial gene by Boubli et al. (2008), as well as the lack of complementary evidences such as cranial or morphometric analyses. Another important question is the proper definition of melanocephalus. The holotype of Cacajao melanocephalus (Humboldt, 1812) is a pet with imprecise origin, but referred as “Exact locality unknown, in the region of the Mission of San Francisco Solano on the Canal Cassiquiare, Amazonas, Venezuela” (Boubli et al. 2008). Boubli et al. (2008) designated a neotype for C. melanocephalus (MN68616; type locality: Serraria, right bank of Rio Negro, Amazonas, Brazil). Surprisingly, Ferrari et al. (2014), disregarding Boubli et al. (2008) designation, selected another neotype for C. melanocephalus (NMNH 406425, type locality: Río Mavaca, 108 km SSE Esmeralda, Amazonas state, Venezuela), claiming that this specimen is closest to the Humboldt’s holotype coloration and geographical origin. Nevertheless, this second designation has no validity (ICZN 1999, Article 75.4). The phylogenetic analysis of Boubli et al. (2008) revealed genetic distances (Tamura-Nei+G) between ayresi and melanocephalus (including sequences of neotype MN68616) ranging from 0.021 to 0.025 while distances between hosomi (= C. m. melanocephalus sensu Ferrari et al. [2014]) and ayresi were conspicuously lower, ranging from 0.003 to 0.006. The hosomi-ayresi range overlaps with the intraspecific distances found for both hosomi (0.000-0.003) and ayresi (0.001-0.004) (Boubli et al. 2008). Furthermore, hosomi and ayresi are sister clades in the Cytb phylogenetic tree and the dichotomy between theses clades is overly shallower when compared to melanocephalus and hosomi + ayresi dichotomy (Fig. 2 in Boubli et al. 2008). The divergence between hosomi and ayresi Cytb sequences is restricted to one C-T transition (Table II in Boubli et al. 2008). In addition, Figueiredo-Ready et al. (2013) found very low Cytb divergence between ayresi and melanocephalus (content not specified), ranging from 0.0004 to 0.007. Thus, to our understanding, (1) there is no genetic divergence that justifies the treatment of hosomi and ayresi as distinct species, and (2) the darker dorsal coloration of first in relation to the later may represent intraspecific geographic variation. Considering these issues, we recommend to treat ayresi Boubli et al. 2008, hosomi Boubli et al. 2008 and melanocephalus [sensu Boubli et al. (2008) and sensu Ferrari et al. (2014)] as Cacajao melanocephalus (Humboldt, 1812) until the light of a more integrative approach, including phylogenetic analyses using mtDNA and nuDNA markers associated to robust morphological analyses (e.g. craniometrics, cranial geometric morphometrics) of a broader sample (including specimens from Venezuela). In relation to C. ouakary, Figueiredo-Ready et al. (2013) found a considerable Cytb divergence between ouakary (including specimens from Solimões river) and melanocephalus (including ayresi) sequences (0.021-0.038). Thus, considering this evidence, it seems appropriate to treat ouakary and melanocephalus as distinct species. Further studies are still needed to clarify the taxonomy of Cacajao melanocephalus species complex.

• 5. Byrne et al. (2016) performed a phylogenetic analysis of Callicebus (sensu Hershkovitz 1988) using a dataset of 20 nuDNA and two mtDNA markers. The authors found deep structuring and proposed nomenclatural changes, erecting genera Cheracebus and Plecturocebus to allocate divergent clades composed of species formerly treated as Callicebus. Byrne et al. (2016) restricted Callicebus Thomas, 1903 to personatus, coimbrai, barbarabrownae, melanochir and nigrifons.

• 6. The genus Plecturocebus was erected by Byrne et al. (2016) to allocate a highly divergent clade of species formerly treated as Callicebus (see remark 5).

• 7. Byrne et al. (2016) considered Callicebus dubius Hershkovitz, 1988 a junior synonym of Callicebus caligatus (Wagner, 1842) (allocated in the new genus Plecturocebus) in view of the monophyletism of caligatus and dubius mtDNA sequences and their very low divergence of nuDNA and concatenated nuDNA and mtDNA sequences. The low divergence between caligatus-dubius is also corroborated by Hoyos et al. (2016) (Cytb) and Carneiro et al. (2016) (Alu elements, 16S, COI, and Cytb). Meanwhile, Serrano-Villavicencio et al. (2016) analyzed the geographic variation in the pelage color of Callicebus [Plecturocebus according to Byrne et al. (2016)] occurring between the Madeira and Purus rivers and considered Callicebus caligatus as a polytypic species with two subspecies: Callicebus caligatus caligatus and Callicebus caligatus dubius. Therefore, herein C. dubius is not considered a full species.

• 8. Plecturocebus grovesi (Boubli et al. 2019) (type locality: the community of Novo Horizonte, left bank of the Rio Teles Pires, municipality of Alta Floresta, Mato Grosso state, Brazil) was described from specimens from a small area of Amazon Forest in northern Mato Grosso state.

• 9. Plecturocebus parecis Gusmão et al. 2019 (type locality: “the Rondon II hydroelectric dam on the middle Rio Comemoração, a tributary of the left margin of the Rio Ji-Paraná, municipality of Pimenta Bueno, Rondônia, Brazil”) was described from specimens collected in western Mato Grosso and eastern Rondonia (Gusmão et al. 2019).

• 10. Byrne et al. (2016), considered Plecturocebus stephennashi Roosmalen, Roosmalen & Mittermeier, 2002 a valid species but without including it in their phylogenetic analysis. Serrano-Villavicencio et al. (2016) based on the pelage color considered stephennashi (as Callicebus) a hybrid form between C. c. caligatus and C. c. dubius. Moreover, both authors raise the question about the origin of stephennashi type series. Herein, we tentatively considered P. stephennashi as valid species, but we recommend additional efforts of new sampling and molecular analyses aiming to clarify the status of this taxon.

• 11. The genus Cheracebus was erected by Byrne et al. (2016) to allocate a highly divergent clade of species formerly treated as Callicebus (see remark 5).

• 12. Serrano-Villavicencio et al. (2019) analyzed the pelage color of Pithecia irrorata species group (sensu Marsh 2014) and considered Pithecia mittermeieri Marsh, 2014, P. pissinattii Marsh, 2014 and P. rylandsi Marsh, 2014 junior synonyms of Pithecia irrorata Gray, 1843, alleging lack of robust diagnoses and poorly defined geographic distributions.

• 13. Pithecia vanzolinii Hershkovitz, 1987 (type locality: “Santa Cruz, Rio Eirú, a east bank (southern) tributary of the Rio Juruá, Amazonas, Brazil.”) was originally described as a subspecies of Pithecia irrorata Gray, 1843 and erected to full species by Marsh (2014) based on highly distinctive pelage characters. The specific status of P. vanzolinii was also corroborated by Serrano-Villavicencio et al. (2019), which analyzed the pelage color of Pithecia irrorata species group (sensu Marsh 2014).

Order Lagomorpha

• 1. Sylvilagus tapetillus Thomas, 1913 was recognized as a distinct species apart from Sylvilagus brasiliensis based on phenotypic, molecular and chromosomal traits (Bonvicino et al. 2015, Ruedas et al. 2017). The current geographic range of S. tapetillus is uncertain. Confirmed records are restricted to the Serra do Mar, Rio de Janeiro state (Bonvicino et al. 2015, Ruedas et al. 2017, Silva et al. 2019).

Order Rodentia

• 1. Brucepattersonius nebulosus Abreu-Júnior, Vilela, Christoff, Valiati & Percequillo, 2019 (type locality: Brazil, São Paulo, Bananal municipality, Estação Ecológica do Bananal) was described from specimens collected in Atlantic Forest localities in the states of São Paulo, Rio de Janeiro and Minas Gerais (Abreu-Júnior & Percequillo 2019).

• 2. Calomys mattevii Gurgel-Filho, Feijó & Langguth, 2015 (type locality: Fazenda Regalito, Rio Santa Maria, 20 km E of Flores de Goiás, Municipality Flores de Goiás, Goiás, Brazil) was considered a junior synonym of Calomys expulsus by Gutiérrez & Marinho-Filho (2017). However, phylogenetic analyses show that specimens of C. mattevii (karyotype 2n= 66/NF= 68) from the Cerrado and Caatinga areas of Goiás, Minas Gerais, Bahia, Piauí, Ceará and Pernambuco states form a distinct monophyletic clade apart from individuals with the same karyotype as C. expulsus (2n = 36/ NF= 66) (Gurgel-Filho et al. 2015, Campos B.A.T.P., unpublished data). Therefore, considering the chromosomal and phylogenetic evidences, we herein considered C. mattevii as valid species.

• 3. Pardinãs et al. (2016) erected the genus Castoria to allocate the fossil species Habrothrix angustidens Winge, 1887 and the living species Akodon serrensis Thomas, 1902, which were considered synonyms. The erection of the new genus was needed considering that Cytb and IRBP sequences of Akodon serrensis fell out of Akodon clade within the Akodontini radiation (D’Elía 2003, D’Elía et al. 2003, Smith & Patton 2007, Ventura et al. 2013, Abreu et al. 2014, Pardiñas et al. 2014)

• 4. Deltamys araucaria (type locality [original]: São Francisco de Paula, Rio Grande do Sul State, Brazil) was described from specimens collected in grassland – Araucaria Forest mosaic (Mixed Ombrophilous Forest, Atlantic Forest biome) in Serra Geral highlands of northeastern Rio Grande do Sul state (Quintela et al. 2017). Pardiñas (2018) restricted the type locality of D. araucaria to “5 km by road W Centro de Pesquisas e Conservação da Natureza Pró-Mata/PUCRS, Rio Grande do Sul State, Brazil”.

• 5. Hylaemys seuanezi (Weksler, Geise & Cerqueira, 1999) (type locality: Fazenda União, Município Casimiro de Abreu, estado do Rio de Janeiro) was considered valid species in the list of Percequillo & Gregorin (2017). Meanwhile, this species has been previously recognized by Percequillo (2015) as a synonym of Hylaeamys laticeps (Lund, 1840) (type locality: “Rio das Velhas’s Floddal”, Lagoa Santa, Minas Gerais) without justifications. The type locality of H. laticeps (Lagoa Santa) is inserted in an area of Cerrado nearly to the transitional zone between this biome and the Atlantic Forest in Minas Gerais state and it is inhabited both by representatives of the rodent fauna typical from Cerrado (e.g. Calomys, Cerradomys, Necromys; Brennand et al. 2013) as well as Atlantic Forest forms (Blarinomys, Delomys, Thaptomys; Ávila-Pires 1960). Although the species limits of Hylaeamys from Brazilian Atlantic Forest is satisfactorily defined through qualitative morphology, craniometrics and karyology (Brennand et al. 2013, considering seuanezi as valid), there is a lack of an integrative analysis of the genus, including the use of molecular markers. In view of this, we considered seuanezi as valid in accordance with Brennand et al. (2013), but we recommend the application of molecular techniques aiming to better clarify the taxonomy of Hylaemys in Brazil.

• 6. Juliomys ximenezi (type locality: Parque Nacional de Aparados da Serra [Aparados da Serra National Park], municipality of Cambará do Sul, Rio Grande do Sul State, Brazil) was described from specimens collected in Araucaria Forest (Mixed Ombrophilous Forest, Atlantic Forest biome) in Serra Geral highlands of northeastern Rio Grande do Sul state (Christoff et al. 2016). Only specimens from type locality are known.

• 7. Hurtado & Pacheco (2017) recognized Neacomys amoenus Thomas, 1903 as a full species, distinguished from Neacomys spinosus (Thomas, 1882) by morphological characters and Cytb sequences. The authors also recognized two subspecies, N. a. amoenus Thomas, 1903 and N. a. carceleni Hershkovitz, 1940. Neacomys spinosus (Thomas, 1882) was restricted to mountain cloud forests of Peruvian Amazonia (Hurtado & Pacheco 2017).

• 8. Neusticomys peruviensis (Musser & Gardner, 1974) was recorded for Parque Nacional de Pacaás Novos, Rondônia state (Percequillo et al. 2017).

• 9. Oecomys bicolor (Tomes, 1860) was considered a species complex based on mitochondrial and nuclear sequences (Suárez-Villota et al. 2017). Four well-supported lineages were recovered within Oecomys bicolor species group. However, the sequence closest to the type locality of O. bicolor [sequence from Peru; type locality of Oecomys bicolor (Tomes, 1860): “Gualaquiza,” Rio Gualaquiza, 885 m, Morona-Santiago, Ecuador (Carleton & Musser 2015)] comprised a clade apart from the Brazilian clades. The name bicolor was herein maintained for the Brazilian Amazon lineages of O. bicolor group until further analyses and formal description of such forms.

• 10. Oecomys catherinae Thomas, 1909 [type locality: “Joinville, Santa Catherina [Santa Catarina], S. Brazil” (Carleton & Musser 2015)] was considered a species complex based on phylogenetic analyses (Suárez-Villota et al. 2017), although no topotypical sequences were employed. One lineage occurs in southeastern Atlantic Forest, one in Cerrado and three in Amazon Forest (Suárez-Villota et al. 2017). The name catherinae was herein maintained but is highly recommended to perform further analyses including sequences from type locality and other localities in southern Atlantic Forest (Santa Catarina and Paraná states).

• 11. Oecomys cleberi Locks, 1981 [type locality: “Fazenda Agua Limpa, da Universidade de Brasília, Distrito Federal, Brasil (Carleton & Musser 2015)] was considered a species complex based on mitochondrial and nuclear sequences (Suárez-Villota et al. 2017), comprising two lineages. One lineage, which includes sequences from the holotype, is distributed in Cerrado of central Brazil. The other occurs in southern Amazon, including Mato Grosso state (Suárez-Villota et al. 2017).

• 12. Oecomys franciscorum was originally described from specimens collected in Argentinean provincies of Formosa and Chaco (type locality: Provincia de Formosa, Departamento de Formosa, Estación de Animales Silvestres Guaycolec, 0.4 km NW of the junction between Ruta Nacional 11 and Riacho Pilagá) (Pardiñas et al. 2016). Cytb sequences of specimens from Brazilian Pantanal (Mato Grosso do Sul state) and treated as Oecomys cf. franciscorum by Pardiñas et al. (2016) grouped with sequences from type specimens in a moderated-supported clade (BPP = 0.96). Following, Suárez-Villota et al. (2017) performed Bayesian phylogenetic analyses using those sequences plus additional specimens from Brazilian Pantanal (including a new locality, Parque Nacional do Pantanal, Mato Grosso state) and found similar relationships and very low K2P distances (0.0 – 1.1%), considering the clade (O. franciscorum – O. cf. franciscorum) a single species.

• 13. Oecomys mamorae (Thomas, 1906) [type locality: type locality “Mosetenes, Upper Mamoré, Yungas, [Cochabamba,] Bolivia.” (Carleton & Musser 2015)] was considered a species complex based on mitochondrial and nuclear sequences, encompassing three lineages (Suárez-Villota et al. 2017). Sequences from Brazil (Pantanal biome, Mato Grosso state), however, were recovered apart from Bolivian lineage, comprising an exclusive clade. The name mamorae is herein maintained until further analyses.

• 14. Oecomys paricola (Thomas, 1904) [type locality: “Igarapé-Assu, near Pará, [Para, Brazil]. Alt. 50 m” (Carleton & Musser 2015)], was considered a species complex based on mitochondrial and nuclear sequences, comprising three lineages (Suárez-Villota et al. 2017). The three lineages are distributed in Brazilian territory, being two in Amazon forest of Para and Mato Grosso states and the other in Cerrado (Suárez-Villota et al. 2017).

• 15. Oecomys roberti (Thomas, 1904) [type locality: “Santa Ana de Chapada, a village situated at an altitude of about 800 m, on the Serra do Chapada, some thirty miles N.E. of Cuyabá [Cuiabá],” Mato Grosso, Brazil (Carleton & Musser 2015)] was revealed as a species complex based on mitochondrial and nuclear sequences, comprising three lineages all occurring in Brazil. One lineage is restricted to Western Amazon, one is widespread from central Amazon to western Cerrado and Pantanal, and the last is restricted to northern Cerrado (Suárez-Villota et al. 2017).

• 16. Oxymycterus itapeby (type locality: Brazil, São Paulo, Itapevi, Transurb district, “Condomínio Vila Verde”) was described from specimens collected in Atlantic Forest-Cerrado transitional areas of São Paulo and Paraná states (Peçanha et al. 2019).

• 17. Rhagomys longilingua Luna & Patterson, 2003 was recorded to Brazilian Amazon in Rondônia state (Hydroelectric Dam Jirau) (Percequillo et al. 2017).

• 18. The type locality of Ctenomys brasiliensis (Blainville, 1826) is indicated as “St Paul, prov. Las Minas” and it was long thought to refer to the state of Minas Gerais, southeastern Brazil, a region apart from the distribution of Ctenomys genus. In view of this, Fernandes et al. (2012) analyzed the holotype of C. brasiliensis through qualitative characters and skull geometric morphometrics. Their results indicated that C. brasiliensis is closely related to C. pearsoni and C. torquatus, respectively distributed in Uruguay and Rio Grande do Sul state and Uruguay. Thus, it is very likely that “Minas” indicated in C. brasiliensis holotype label could be related to the municipality of Minas, the capital of Lavalleja department, Uruguay, a region within the distribution of C. pearsoni. Considering this evidences, we did not consider C. brasiliensis as occurring in Brazil.

• 19. Agouti silvagarciae Van Roosmalen et Van Hoof, 2015 was described from two specimens, one collected by hunters at the left bank of Aripuanã river, and the other vouchered in Museu Paraense Emílio Goeldi, without provenance. Acording to Van Roosmalen (2015), A. silvagarciae differs from C. paca by overall large size. The author also states that “One complete mitochondrial Dloop and two nuclear SINE PRE-1 DNA sequences of Silva Garcia’s giant paca were carried out and compared with Genbank sequences of the sympatric common paca (A. paca). The results (15.5% difference between species) clearly support the distinction into valid species.” However, the author makes no distinction about the distance found for each marker, and no reference to the methods used for phylogenetic inference, nodal supports, and the C. paca sequences used in the analysis was added. In view of the lack of such crucial information, we herein did not consider A. silvagarciae as valid.

• 20. Dasyprocta aurea Cope, 1889, Dasyprocta catrinae Thomas, 1917 and Dasyprocta nigriclunis Osgood, 1916 were considered by Iack-Ximenes (2019) as valid species. Dasyprocta aurea and D. catrinae were considered synonyms of Dasyprocta azarae Lichtenstein, 1823 while D. nigriclunis is referred as a synonym of D. prymnolopha Wagler, 1831 by Patton & Emmons (2015a).

• 21. Phyllomys centralis (type locality: Fazenda Água Limpa (15°57’4.42’S, 47°57’48.85’W), APA Gama Cabeça de Veado, Distrito Federal, Brazil, elevation of 1,100 m) was described from specimens collected in Cerrado of Distrito Federal state and Atlantic Forest-Cerrado transitional area in Minas Gerais state (Machado et al. 2018).

• 22. Trinomys panema (Moojen, 1948) was considered a synonym of Trynomys gratiosus (Moojen, 1948) by Patton & Emmons (2015b) and herein is not considered as full species.

• 23. Coendou baturitensis Feijó & Langguth, 2013 (type locality: Community Sitio Barreiros, municipality of Aratuba, Baturite Range, Ceará, Brazil) was considered a junior synonym of Coendou prehenselis by Voss (2015) and Gutiérrez & Marinho-Filho (2017). Recent morphology and Cytb phylogenetic analyses revealed that individuals of C. baturitensis cluster apart from C. prehenselis and shows several diagnostic phenotypic traits (Menezes F.H., unpublished data). Therefore, we herein considered C. baturitensis a valid species.

• 24. Coendou ichillus Voss & da Silva, 2001 was reported in Brazil based on one individual collected in the margins of Rio Japurá, Limoeiro, Amazonas state (Menezes et al. 2020).

• 25. Hadrosciurus ignitus (Gray, 1867) was previously considered as a subspecies of Notosciurus pucheranii (Fitzinger, 1867). Abreu-Júnior et al. (2020), based on mitogenomic phylogeny, considered it as a full species and part of the genus Hadrosciurus Allen, 1915. In Brazil, H. ignitus is recorded in the states of Acre and Amazonas.

Order Chiroptera

• 1. Eumops chimaera (type locality: Parque Estadual do Rio Doce [PERD], municipality of Marliéria, state of Minas Gerais, Brazil) was described from specimens collected in the Atlantic Forest of Minas Gerais state and the Bosque Chiquitano of Bolivia (Gregorin et al. 2016). Eumops chimaera diverges from other Eumops species by Cytb sequences and external and craniodental morphology (Gregorin et al. 2016).

• 2. Pteronotus (Phyllodia) alitonus Pavan, Bobrowiec & Percequillo, 2018 was described from specimens from Suriname, French Guiana, and the Brazilian Amazon, previously referred as Pteronotus sp. 1 (Pavan & Marroig 2016), Pteronotus sp. 3 (Clare et al. 2013), Pteronotus rubiginosus and P. parnellii (see Pavan et al. 2018). Pteronotus (Phyllodia) alitonus differs from other Pteronotus species by morphological traits, echolocation calls and COI sequences (Pavan et al. 2018). Pteronotus parnellii Gray, 1843 (type locality: Jamaica) was excluded from the Brazilian fauna because none of the Brazilian specimens analyzed shared haplotypes with the Jamaican lineage (see Thoisy et al. 2014, López-Wilchis et al. 2016).

• 3. Pteronotus Gray, 1838 was organized in three subgenera (Pteronotus, Phyllodia, Chilonycteris) by Smith (1972) based on a robust craniometric dataset. However, molecular analyses performed by Pavan & Marroig (2016) using mitochondrial and nuclear markers (COI, Cytb, Dby, RAG2, STAT5A) revealed Chilonycteris as an artificial group and suggested the erection of a new subgenus for Pteronotus personatus (Wagner, 1843). Since there is still no available name, we kept a binomial classification for P. personatus.

• 4. Chiroderma salvini Dobson, 1878 was recorded for Brazil by Rocha et al. (2016) based on two specimens from Rondônia and Mato Grosso states. However, analysing a large sample of Chiroderma, Brandão et al. (2019) argued that both specimens reported by Rocha et al. (2016) are actually C. villosum. Therefore, we did not include C. salvini as part of the Brazilian fauna.

• 5. Garbino et al. (in press) revised the genus Chiroderma using morphological and molecular datasets and recognized Chiroterma vizottoi Taddei & Lim, 2010 as a subspecies of Chiroderma doriae Thomas, 1891. Therefore, C. vizottoi is not included in the present list.

• 6. The genus Gardnerycteris Hurtado & Pacheco, 2014 was erected to allocate two species previously included in genus Mimon, which was revealed as polyphyletic from a phylogenetic analysis of 91 morphological characters (Hurtado & Pacheco 2014). Later, the monophyly of Gardnerycteris and its contents were sustained by molecular data (Cytb, COI, RAG2) (Hurtado & D’Elía 2018). Gardnerycteris currently comprises three valid species (G. koepckeae [type species], G. crenulatum and G. keenani) and G. crenulatum is the single species occurring in Brazil (Hurtado & D’Elía 2018).

• 7. Hsunycteris pattoni (Woodman & Timm, 2006), originally described as Lonchophylla pattoni (type locality: Reserva Cusco Amazónico, north bank of the Río Madre de Dios; 14 km east of Puerto Maldonado; Tambopata Province; Madre de Dios Department; southeastern Peru), was assigned to Brazilian Amazon in the states of Amazonas and Pará (Velazco et al. 2017).

• 8. Lonchophylla inexpectata Moratelli & Dias, 2015 (type locality: Barra, Bahia, Brazil) was described from specimens previously identified as Lonchophylla mordax Thomas, 1903. Lonchophylla inexpectata is distinguishable from other Lonchophylla species by the fur color, cranial size and dental morphological (Moratelli & Dias 2015).

• 9. Micronycteris brosseti Simmons & Voss, 1998 (type locality: Paracou, French Guiana) was not considered in the present list. The referred material of M. brosseti from Brazil includes only one specimen from “Rio Juquiá, Barra, São Paulo”, deposited in the Field Museum of Natural History, Chicago (FMNH 92997) (Simmons & Voss 1998). Garbino (2016) disagrees with this identification, taking into account a personal communication from Ricardo Moratelli, who alleged that FMNH 92997 could not be conclusively identified as M. brosseti. Garbino (2016) also called attention to the over 3,000 km gap between the São Paulo record and the type locality of M. brosseti. Considering these issues, we opted for the exclusion of M. brosseti until further clarification.

• 10. Sturnira giannae Velazco & Patterson, 2019 (type locality: Paracou (5°17’N, 53°55’W, 210 m), near Sinnamary, Cayenne, French Guiana) was described from specimens collected in Guianas, Venezuela, Ecuador, Peru, Bolívia and the Brazilian states of Amazonas and Pará (Velazco & Patterson 2019).

• 11. Tonatia maresi Williams, Willig & Reid, 1995 (type locality: Blanchisseuse, Trinidad and Tobago) was originally described as a subspecies of Tonatia saurophila Koopman & Williams, 1951. Basantes et al. (2020) elevated maresi and T. saurophila bakeri Williams, Willig & Reid, 1995 to species level based on morphological and genetic (Cytb and nuclear exon RAG2) differences. Tonatia currently comprises three valid species (T. saurophila [considered extinct and restricted to Jamaica], T. bakeri, and T. maresi), and only T. maresi is recorded in Brazil (Basantes et al. 2020).

• 12. Histiotus diaphanopterus Feijó, Rocha & Althoff, 2015 (type locality: Boqueirão do Onça, village of São Pedro do Lago, municipality of Sento Sé, state of Bahia, Brazil) was described from specimens collected in Caatinga and Cerrado of Brazil (Bahia, Ceará, Paraíba and Maranhão states) and Bosque Chiquitano of Bolivia.

Order Carnivora

• 1. The name Conepatus semistriatus (Boddaert, 1785) has been historically applied for the form occurring in Cerrado and Caatinga of Brazil (e.g. Cavalcanti et al. 2013, 2014, Dias 2017, Tomas et al. 2017). However, phylogenetic analyses using mtDNA markers (Cytb, COI, CR) recovered sequences from a specimen from Mexico (type locality of C. semistriatus) in a clade apart from sequences of specimens from South America, including individuals from Brazilian Cerrado-Caatinga (states of Goiás, Minas Gerais and Piauí) (Schiaffini et al. 2013). This aspect was observed by Feijó & Langguth (2013), who stated that C. amazonicus (Lichtenstein, 1838) is the name applicable to Conepatus from Cerrado and Caatinga in Brazil. Herein we follow this new classification.

• 2. Kitchener et al. (2017) considered the jaguarundi as belonging to genus Herpailurus Severtzov, 1858, only stating “As used here, this is a monotypic genus, but it may be included within Puma”. Herein, Herpailurus yagouaroundi is used instead of Puma yagouaroundi.

• 3. Nascimento et al. (2020), using an integrative taxonomic approach, recognized five species of pampas cats (Leopardus braccatus, L. colocola, L. garleppi, L. munoai, and L. pajeros). These five species show distinct skull and skin traits, inhabit distinct climatic niches and are supported by phylogeny and molecular species delimitation. In Brazil, two species of pampas cats are present. Leopardus braccatus (Cope, 1889) (type locality: Chapada dos Guimarães, Mato Grosso, Brazil) is distributed in central Brazil (from Cerrado of south-western Piauí to Pantanal of Mato Grosso do Sul), Paraguay, and open areas of Bolivia and northern Argentina.

• 4. Leopardus emiliae (Thomas, 1914) (type locality: “Ipu, Ceará, N.E. Brazil. Alt. 300 m”) has long been considered a junior synonym of Leopardus tigrinus (Wozencraft 2005). Nascimento & Feijó (2017) resurrected L. emiliae as a full species, distinguishable from L. tigrinus (Schreber, 1775) and L. guttulus (Hensel, 1872) by pelage color and cranial differences.

• 5. Leopardus munoai (Ximénez, 1961) (type locality: ‘Arroyo Perdido, Departamento de Soriano’, Uruguay) is considered as a valid species by Nascimento et al. (2020) (see remark 3). It occurs in the pampas of Rio Grande do Sul Brazilian state, Uruguay and northeastern Argentina (Nascimento et al. 2020).

Order Perissodactyla

• 1.Tapirus kabomani was described from specimens collected in Rondônia state, southern Brazilian Amazonia (Cozzuol et al. 2013). The authors concluded that the new tapir differs from T. terrestris by external and cranial external morphology and molecular characters (Cytb, COI, COII). Meanwhile, Voss et al. (2014b) discussed the validity of T. kabomani claiming the very low molecular divergence from T. terrestris (1.3%) and the inconsistence between molecular and morphological characters, among other aspects. Herein we tentatively considered T. kabomani as a valid species, but in agreement with Voss et al. (2014b), we believe that further molecular analysis including nuclear markers could provide a better view on T. kabomani taxonomic status.

Order Artiodactyla

• 1. The name Cetartiodactyla Montgelard, Catzeflis & Douzery, 1997 was proposed as a “solution” for the paraphyletic condition of Artiodactyla in relation to Cetacea Brisson, 1762. The International Code of Zoological Nomenclature (ICZN) does not regulate order names, and considering that Cetacea is a clade within Artiodactyla (Montgelard et al. 1997), there is no restriction for the usage of Order Artiodactyla as a conservative name. Therefore, herein we adopted Order Artiodactyla.

• 2. Mazama tienhoveni van Roosmalen & van Hooft, 2015 was described from two skins, one skull and mandible, and one spike in possession of hunters from Tucunaré village, lower Aripuanã River, Central Amazon. In our point of view, the new species presents some inconsistencies, including: 1) The holotype is poorly defined. According to the author the “holotypus” MR204 consists of “complete head with partly damaged mandible, adult female, on May 12, 2006 killed for food by a local hunter along the left bank of the Rio Aripuanã near the settlement of Tucunaré, skull, spike and skin”. The author mentions “two skins” in “Examined material” section but also mentions that “No paratypes have been collected thus far” in “Variability” section. In “Remarks” section, Van Roosmalen (2015) also included: “The males of M. tienhoveni n. sp. do not have the distinct crest of hairs on the forehead as M. nemorivaga has, neither do the males of Mazama americana”. These information raised the following questions: Does this material, assigned as “holotypus MR204” comprise more than one individual? This doubt arose taking into account the references of: a) two skins; b) a spike as part of the material designated as an “adult female”. 2) Van Roosmalen (2015) performed a dated linearized minimum-evolution tree using a very small fragment consisted of 233 bp of Cytb. We believe that the small fragment used for phylogenetic inferences is little informative. The resultant phylogenetic tree, in turn, showed a basal polytomy, with unsolved relationships (Mazama genus was recovered as polyphyletic) and undetermined or weak supports of clades. In summary, the molecular dataset used by Roosmalen is insufficient for a phylogenetic inference within Mazama and the support of a specific-level divergent form. 3) Van Roosmalen (2015) made comparisons between M. tienhoveni and the other Amazonic representatives of Mazama genus, M. americana and M. nemorivaga. Those comparisons were based on pelage coloration, morphometrics (spike size, 16 cranial measurements, tail length, ear length, hindfoot length, head-body length, cranial) and body weight. However, in our understanding, the material comprised by two skins, a skull and mandible and one spike is insufficient for a solid comparison of high variable characters such as body and skull measurements and pelage color. Furthermore, head-body length and body mass were not directly obtained but were assumed to be intermediate between M. americana and M. nemorivaga, based on hunter’s information. Craniometric dataset of other species were also very limited, comprising 11 specimens of M. nemorivaga and five specimens of M. americana, without any distinction on age classes or sex. Moreover, only means of M. americana and M. nemorivaga cranial measurements were presented while ranges were not disclosed. Considering these aspects, we herein did not consider Mazama tienhoveni a valid species due to the lack of genetic and morphological support.

• 3. Tursiops gephyreus Lahille, 1908 was revalidated for Tursiops forms occurring in estuaries of Brazilian states of Paraná, Santa Catarina and Rio Grande do Sul state, Uruguay and Argentina), based on morphometrics and qualitative morphological characters (Wickert et al. 2016). According to these authors, Tursiops truncatus (Montagu, 1821) also occurs in Brazilian coast, along all coastline, but more associated with open waters. Meanwhile, posterior molecular analyses (CR, nuclear microsatellites) revealed few support for the designation of two species of bottlenose dolphins occurring in Brazilian coast (Oliveira et al. 2019). First, “truncatus haplogroup” and “gephyreus haplogroup” are separated by a single mutational step (see Fig. 2 in Oliveira et al. 2019). Second, there was a lack of complete structuring between the genotypes of the morphotypes attributed to gyphereus and truncatus (Oliveira et al. 2019). In view of this, Oliveira et al. (2019) recommended further integrative analyses on a broader geographical sample for a formal decision on the taxonomic status of T. gephyreus. In agreement, we considered only T. truncatus as valid.

• 4. Inia araguaiaensis Hrbek, Farias, Dutra & da Silva, 2014 (type locality: near the entrance of Lake Jurumirim, Araguaia River, state of Goiás) was described from three specimen collected in Araguaia River, state of Goiás. Inia araguaiaensis differs from other Inia species by morphological and molecular (Cytb, COI, nuDNA microsatellite markers) characters. The status of I. araguaiaensis, however, was argued by The Society for Marine Mammalogy (2019), alleging sampling gaps which would leave doubts about the genetic difference found by Hrbek et al. (2014) represents a specific level divergence or an effect of isolation by distance in a specific clade. The Society for Marine Mammalogy (2019) also argues about the morphological analysis based on only two specimens of I. araguaiaensis and nine specimens of I. geoffrensis, which could not account possible effects of sexual dimorphism between the species. Herein we recognized I. araguaiaensis in view of the clear marked genetic structure (see Hrbek et al. 2014), but we also recognize that morphological analyzes on a broader sample could give more support to the specific status of I. araguaiaensis.

Introduced (non-native) species

To date, eight species of mammals are known to have been introduced to Brazil:

Small rodents (Mus musculus Linnaeus, 1758, Rattus rattus (Linnaeus, 1758) and Rattus norvegicus (Berkenhout, 1769)): the three murid rodents were unintentionally introduced in South America in the sixteenth century, during the European colonization of the continent (Pimentel 2011). The three species, originally from Eurasia, spread into Brazil and now occur in all biomes and states of the country, in both conserved and disturbed areas, including protected areas (Rosa et al. 2017a).

Lepus europaeus Pallas, 1778: in the late nineteenth century, the European hare L. europaeus, native in Eurasia, arrived in Argentina and Chile to serve as hunting. It spreaded to Southern Brazil in the 1950s by natural dispersal across the border with Uruguay and through deliberate introductions (Grigera & Rapoport 1983, Costa & Fernandes 2010). Nowadays, L. europaeus occupies mainly pastures, agricultural areas and forest edges of Pampa, Cerrado and Atlantic Forest biomes in the Brazilian states of Rio Grande do Sul, Santa Catarina, Paraná, São Paulo, Minas Gerais, Goiás and Bahia.

Sus scrofa Linnaeus, 1758: the wild boar S. scrofa is native in Eurasia and Africa and was first introduced to South America in the sixteenth century for meat consumption by explorers and settlers. By the late twentieth century, S. scrofa had reached Brazil from Uruguay and in the beginning of the twenty-first century several independent introductions were made throughout Brazil for meat production and hunting (Oliveira C.H.S., unpublished data). There are currently both voluntary introductions for hunting purposes and involuntary introductions of animals escaping from illegal domestic breeding (Oliveira C.H.S., unpublished data). Nowadays, S. scrofa occupies both conserved and disturbed areas throughout Brazil, including protected areas, of all biomes in the states of Rio Grande do Sul, Santa Catarina, Paraná, São Paulo, Minas Gerais, Espírito Santo, Rio de Janeiro, Goiás, Tocantins, Mato Grosso, Mato Grosso do Sul, Bahia, Ceará, Pará, Maranhão and Rondônia.

Axis axis (Erxleben, 1777): original from Asia, the chital A. axis was first recorded in 2010 at Espinilho State Park, Rio Grande do Sul state, in Southern Brazil, probably coming from Argentina where the species was introduced for hunting (Sponchiado et al. 2011, Rosa et al. 2017a). Since then, A. axis have been reported in several municipalities of Rio Grande do Sul (Rosa et al. 2017a), occupying grasslands of Pampa biome where feral populations may have established.

Bubalus bubalis (Linnaeus, 1758): original from Asia, the water buffalo B. bubalis was intentionally introduced to Brazil in the nineteenth century for food and labor animal (Rosa et al. 2017a). Nowadays B. bubalis has scattered populations in both conserved and disturbed areas, including protected areas, in the Pampa, Atlantic Forest, Pantanal, Cerrado and Amazon biomes in the states of Rio Grande do Sul, Santa Catarina, Paraná, São Paulo, Rio de Janeiro, Goiás, Minas Gerais, Mato Grosso, Mato Grosso do Sul, Pará, Amapá, Rondônia, Bahia, Tocantins, Maranhão and Piauí (Rosa et al. 2017a).

Rusa unicolor (Kerr, 1792): the sambar R. unicolor is original from Asia and was intentionally introduced during the twentieth century into non fenced areas of Atlantic Forest, likely for sport hunting (Pimentel 2011). Nowadays three isolated populations of R. unicolor are known, one in Paraná state and two in São Paulo state (Pimentel 2011, Rosa et al. 2017a).

Native displaced species

Seven species have been translocated from their natural habitats in Brazil to other locations inside the country:

Callithrix sp. (Callithrix jacchus (Linnaeus, 1758), Callithrix geoffroyi (Humboldt, 1812), Callithrix penicillata (É. Geoffroy St.-Hilaire, 1812) and their hybrids): Three species of marmosets from Callithrix genus have been introduced through Brazil, including islands and protected areas, as a result of pet escapes and misguided releases of confiscated pet animals (Pimentel 2011). Callithrix jacchus naturally occurs in the scrub forests of the Atlantic Forest in Northeast Brazil (Bezerra et al. 2018). Invasive populations of the C. jacchus have been reported in several Atlantic Forest regions in states of Bahia, Minas Gerais, Rio de Janeiro, São Paulo, Espírito Santo, Paraná and Santa Catarina (Rosa et al. 2017a). Callithrix penicillata has a wide native distribution in Brazil, occurring in gallery forests, dry forests, and forest patches in the Cerrado, Caatinga and Atlantic Forest (Bicca-Marques et al. 2018). Invasions of C. penicillata have been reported in Atlantic Forest habitats of states of Rio Grande do Sul, Santa Catarina, Paraná, Minas Gerais, São Paulo, Rio de Janeiro, Bahia and Paraíba. Callithrix geoffroyi originally occurs in the Atlantic Forest of Minas Gerais, Bahia and Espírito Santo (Rylands & Mendes 2018) and is known to have been introduced in southern Brazilian Atlantic Forest, into Florianópolis Island in the state of Santa Catarina. The three species hybridize with each other and with Callithrix flaviceps, Callithrix kuhlli and Callithrix aurita. Hybrids were registered in the states of Santa Catarina, Paraná, Espírito Santo, Rio de Janeiro, São Paulo, Minas Gerais, Pernambuco and Bahia (Rosa et al. 2017a, Aximoff et al. 2020).

Saimiri sciureus (Linnaeus, 1758): the common squirrel monkey S. sciureus is a small Amazonian primate and was intentionally introduced to the Atlantic Forest. The first record of S. sciureus outside its natural range was in 1987 at the Saltinho Biological Reserve, Pernambuco, during a pet release operation of individuals apprehended from illegal fauna trade (Camarotti et al. 2015). Currently, established populations of S. sciureus occur in forest fragments of Atlantic Forest in the states of Pernambuco, Rio de Janeiro and Alagoas. The species has potential to expand its distribution into non-native environments due illegal pet release (Rosa et al. 2017b).

Leontopithecus chrysomelas (Kuhl, 1820): the golden-head lion tamarin L. chrysomelas is an endemic and threatened species of the Atlantic Forest of Bahia state (Coimbra-Filho & Mittermeier 1973, Kierulff et al. 2008). In the 1990s, individuals of L. chrysomelas were accidentally introduced by a private collector into an urban Atlantic Forest remnant in the state of Rio de Janeiro, where established an invasive population (Kierulff et al. 2012, Molina et al. 2017). Recently, some individuals of L. chrysomelas were also seen in forest fragments of Atlantic Forest in Camaragibe municipality, Pernambuco state (Rosa et al. In Press).

Myocastor coypus (Molina, 1782): the nutria M. coypus is a large rodent native from open areas of southern South America, including the south of Brazil (Ojeda et al. 2016). In Brazil M. coypus was introduced by breeders for purposes of use of their skin (Bueno 2013). The species established non-native populations in states of São Paulo, Rio de Janeiro and Minas Gerais (Rosa et al. In Press).

Kerodon rupestris (Wied Neuwied, 1820): the rock cavy K. rupestris is a rodent native to the Brazilian Caatinga biome. In 1967, the species was introduced to Fernando de Noronha Archipelago by military personnel as hunting game, and an established population occurs until today (Pimentel 2011, Rosa et al. 2017a). No other non-native populations of K. rupestris are known to occur in Brazil.

Discussion

Brazilian mammalogy is passing through an accelerated period of investigative taxonomy. The first compilation of Brazilian mammals (Fonseca et al. 1996) listed 524 species. Ten years later, Reis et al. (2006) indicated the occurrence of 652 species to the country. This number was increased to 694 species after five years (Reis et al. 2011). The following compilation, the annotated checklist of Paglia et al. (2012), published only one year later, listed a total of 701 species. After five years, the last list of mammals occurring in Brazil (Percequillo & Gregorin 2017) presented a total of 721 species. During the short period of two years between the publication of Percequillo & Gregorin (2017) list and the annotated checklist herein presented, 15 new taxa were described (Miranda et al. 2018, Pavan et al. 2017, Quintela et al. 2017, Machado et al. 2018, Pavan et al. 2018, Voss et al. 2018, Boubli et al. 2019, Pavan 2019, Peçanha et al. 2019, Velazco & Patterson 2019, Costa-Araújo et al. 2019, Abreu-Júnior & Percequillo 2019), most of them supported by genetic analyses. This, added to synonimizations (e.g. Byrne et al. 2016, Serrano-Villavicencio et al. 2019) and to the new records for the country (e.g. Bezerra et al. 2019, Percequillo et al. 2017, Menezes et al. 2020), resulted in 751 species herein listed. Notwithstanding, our checklist represents the entire mammalian diversity so far known for the country, the available evidence pointed out a still underestimated diversity. Based on recent taxonomic revisions, some of the taxa currently listed clearly represent species complex that invites further studies (e.g. Oecomys; Suárez-Villota et al. 2017). In addition, several lineages divergent at specific level identified through molecular phylogenetic analyses still await formal description. These lineages comprise new forms of rodent genera Deltamys (Quintela et al. 2017), Holochilus (D’Elía et al. 2015), and Phyllomys (Machado et al. 2018), a new form of long-nosed armadillo (Dasypus) from Guiana Shield region (Feijó et al. 2019), among others. Thus, the new era of integrative taxonomic studies combining morphology and molecular tools is unveiling an overlooked diversity and driving a new period in Brazilian mammalogy. Hence, a considerable increment in the Brazilian mammalian diversity is expected. The increase in the number of researchers dedicated to systematics and taxonomy of mammals, together with greater field survey efforts in new regions, are also important factors that have contributed to advancement on the knowledge of mammalian diversity in Brazil.

The mammalian richness in Brazil is considerably higher than those reported in surrounding countries, including Argentina (409 spp.; Teta et al. 2018), Bolivia (406 spp; Aguirre et al. 2019), Paraguay (184 spp.; de la Sancha et al. 2017, 2019), Peru (508 spp; Pacheco et al. 2009) and Uruguay (73 spp; Queirolo 2016). This higher diversity may be a reflex of the larger area and greater diversity of environments in Brazil in relation to neighboring countries.

The negative effects of introduced (non-native or native) mammalian species are far known (see Rosa et al. 2017a). Reis et al. (2006, 2011) also concerned on listing the non-native species established in natural environments in Brazil and in that time this species were restricted to the murid rodents M. musculus, R. novergicus and R. rattus, the European hare L. europaeus, the wild boar S. scrofa and the water buffalo B. bubalis. Since them, the cervids Axis axis and Rusa unicolor were detected in the wild respectively in Atlantic Forest-Cerrado transitional area and Pampa environments, raising to eight the number of non-native mammals introduced to the country. Another very relevant issue is the occurrence of native displaced species, comprising five species of primates and two species of hystricomorph rodents. The potential impacts of both native and non-native species, as well as the recommendations for management and policy, are discussed in Rosa et al. (2017a).

Around 10% (80 species) of the Brazilian mammalian fauna is threatened in a global scale. However, the Brazilian Red Book of Threatened Species of Fauna (ICMBio 2018) listed a total of 110 taxa (15% of the total species), which implies that many species considered regionaly threatened (e.g. Thylamys macrurus, T. velutinus, Ozotoceros bezoarticus, Leopardus geoffroyi, L. guttulus, Puma concolor) are not considered globally threatened. This situation is especially notable for order Carnivora considering that only two from the 12 species included in the Brazilian list (ICMBio 2018) were considered as threatened by IUCN (2019). On the other hand, most of the primate species considered regionally threatened were also evaluated as globally threatened. Together with these two orders, rodents also contribute substantially for both Brazilian and global lists. Most of the primate and rodent threatened species are endemic to Brazil. In general, the Brazilian mastofauna has an impressive endemism rate, comprising more than a quarter of the species accounted for the country. Around 22% of the endemic species (50 species) are threatened and a similar percentage has Data Deficient for conservation according to IUCN (2019). Therefore, endemic species comprise the greatest part of the threatened mammalian fauna in Brazil (62%). Many of these species are distributed mainly in non-protected areas as well as the non-endemic threatened taxa. The loss, fragmentation and decharacterization of the natural environments represent the greatest threats to the mammalian fauna in Brazil (Costa et al. 2005, ICMBio 2018), and outside the protected areas, populations are extremely susceptible to these deleterious factors. Thus, the creation of new conservation units and the expansion of the established protected areas are crucial for mammalian conservation in Brazil.

The near future of mammalogy in Brazil is a quite delicate matter. On the one hand, we have an enormous potential for the discovery of new species and a wide field of research for many systematic, evolutionary, ecology, phylogeography, population genetics open questions. On the other hand, we have an unfavorable scenario for the biological conservation in the country. The current Brazilian Forest Code (established in 2012) is very ineffective for the maintenance of the areas covered by native vegetation, and is especially damaging to Atlantic Forest and Cerrado biomes, where only 20% of the natural areas is required as preserved within private properties. Atlantic Forest and Cerrado are recognized biodiversity hotspots (Myers et al. 2000) and have a crucial role in mammalian conservation in Brazil (Costa et al. 2005). Moreover, the new Brazilian government is absolutely antagonistic to the scientific development and biological conservation. The Ministry of Environment has drastically weakened on its crucial role of regularization of rural properties and control of the protected areas. Additionally, there has been a dramatic budget reduction in scientific research in recent years. In summary, the current panorama of uncertainty and instability of Brazilian science and weaken environmental rules are not consistent with the status of the country as the world’s highest biodiversity, which includes an impressive mammalian fauna.

ACKNOWLEGMENTS

We are grateful to Guilherme Garbino, Fabrício Bertuol, Ulyses Pardiñas, and two anonymous reviewers for the meaningful suggestions. FMQ is supported by Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES, fellowship process PNPD201321692-42004012013P2). AF is supported by Chinese Academy of Sciences President´s International Fellowship Initiative (Grant N°2018PB0040). CAR is supported by Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq, fellowship process 300921/2019-5).

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APPENDIX

List of mammal species occurring in Brazil and their conservation status based on the Brazilian (ICMBio 2018) and international lists of threatened species (IUCN 2019). Acronyms: CR = critically endangered, DD = Deficient Data, EN = endangered, EX = presumable extinct, LC = least concern, NT = near threatened, PE = pending (re)evaluation, VU = vulnerable. Numbers between parenthesis, after authorship and date of description, indicate remarks included in the main text (Results section), separated by order. Asteriscs (*) indicate the species endemic to the country.

Taxon	Conservation Status
Class Mammalia Linnaeus, 1758	IUCN	ICMBio
Infraclass Marsupialia
Order Didelphimorphia Gill, 1872
Family Didelphidae Gray, 1821
Caluromys Allen, 1900
Caluromys philander (Linnaeus, 1758)	LC
Caluromys lanatus (Olfers, 1818)	LC
Caluromysiops Sanborn, 1951
Caluromysiops irrupta Sanborn, 1951	LC	CR
Chironectes Illiger, 1811
Chironectes minimus (Zimmermann, 1780)	LC
Cryptonanus Voss, Lunde & Jansa, 2005
Cryptonanus agricolai (Moojen, 1943) *	DD
Cryptonanus chacoensis (Tate, 1931)	LC
Cryptonanus guahybae (Tate, 1931) *	DD
Didelphis Linnaeus, 1758
Didelphis albiventris Lund, 1840	LC
Didelphis aurita Wied-Neuwied, 1826	LC
Didelphis imperfecta Mondolfi & Pérez-Hernández, 1984	LC
Didelphis marsupialis Linnaeus, 1758	LC
Glironia Thomas, 1912
Glironia venusta Thomas, 1912	LC
Gracilinanus Gardner & Creighton, 1989
Gracilinanus agilis (Burmeister, 1854)	LC
Gracilinanus emiliae (Thomas, 1909)	DD
Gracilinanus microtarsus (Wagner, 1842)	LC
Gracilinanus peruanus (Tate, 1931) (1)	PE
Hyladelphys Voss, Lunde & Simmons, 2001
Hyladelphys kalinowskii (Hershkovitz, 1992)	LC
Lutreolina Thomas, 1910
Lutreolina crassicaudata (Desmarest, 1804)	LC
Marmosa Gray, 1821
Marmosa (Marmosa) murina (Linnaeus, 1758)	LC
Marmosa (Micoureus) constantiae (Thomas, 1904)	LC
Marmosa (Micoureus) demerarae (Thomas, 1905)	LC
Marmosa (Micoureus) paraguayana Tate, 1931	LC
Marmosa (Micoureus) phaea Thomas, 1899 (2)	VU
Marmosa (Micoureus) rapposa Thomas, 1899 (3)	PE
Marmosa (Micoureus) regina (Thomas, 1898)	LC
Marmosa (Micoureus) rutteri Thomas, 1924 (4)	PE
Marmosa (Stegomarmosa) lepida (Thomas, 1888) (5)	LC
Marmosops Matschie, 1916 (6,7)
Marmosops (Marmosops) incanus (Lund, 1840) *	LC
Marmosops (Marmosops) neblina Gardner, 1989	LC
Marmosops (Marmosops) noctivagus (Tschudi, 1845)	LC
Marmosops (Marmosops) ocellatus (Tate, 1931)	LC
Marmosops (Marmosops) paulensis (Tate, 1931) *	LC	VU
Marmosops (Sciophanes) bishopi (Pine, 1981)	LC
Marmosops (Sciophanes) parvidens (Tate, 1931)	LC
Marmosops (Sciophanes) pinheiroi (Pine, 1981)	LC
Metachirus Burmeister, 1854
Metachirus myosuros (Temminck, 1824) (8)	PE
Metachirus nudicaudatus (Desmarest, 1817)	PE
Monodelphis Burnett, 1830 (9,10)
Monodelphis (Microdelphys) americana (Müller, 1776) *	LC
Monodelphis (Microdelphys) scalops (Thomas, 1888)	LC
Monodelphis (Microdelphys) iheringi (Thomas, 1888) *	DD
Monodelphis (Monodelphis) arlindoi Pavan, Rossi & Schneider, 2012	PE
Monodelphis (Monodelphis) brevicaudata (Erxleben, 1777)	LC
Monodelphis (Monodelphiops) dimidiata (Wagner, 1847)	LC
Monodelphis (Monodelphis) domestica (Wagner, 1842)	LC
Monodelphis (Monodelphis) glirina (Wagner, 1842)	LC
Monodelphis (Monodelphis) touan (Shaw, 1800)	PE
Monodelphis (Monodelphis) vossi Pavan, 2019 (11) *	PE
Monodelphis (Monodelphiops) unistriata (Wagner, 1842)	CR
Monodelphis (Mygalodelphys) handleyi Solari, 2007 (12)	DD
Monodelphis (Mygalodelphys) kunsi Pine, 1975	LC
Monodelphis (Mygalodelphys) pinocchio Pavan, 2015 *	PE
Monodelphis (Mygalodelphys) saci Pavan, Mendes-Oliveira & Voss, 2017 (13) *	PE
Monodelphis (Pyrodelphys) emiliae (Thomas, 1912)	LC
PhilanderBrisson, 1762
Philander andersoni (Osgood, 1913)	LC
Philander canus (Osgood, 1913) (14)	PE
Philander quica (Temminck, 1824) (15)	PE
Philander mcilhennyi Gardner & Patton, 1972	LC
Philander opossum (Linnaeus, 1758)	LC
Philander pebas Voss, Díaz-Nieto & Jansa, 2018 (16)	PE
Thylamys Gray, 1843
Thylamys karimii (Petter, 1968) *	VU
Thylamys macrurus (Olfers, 1818)	NT	EN
Thylamys velutinus (Wagner, 1842) *	NT	VU
Infraclass Placentalia
Superordem Afrotheria
Order Sirenia Illiger, 1811
Family Trichechidae Gill, 1872
Trichechus Linnaeus, 1758 (1)
Trichechus inunguis (Natterer, 1883)	VU	VU
Trichechus manatus Linnaeus, 1758	VU	EN
Superorder Xenarthra
Order Cingulata Illiger, 1821
Family Dasypodidae Gray, 1821
Dasypus Linnaeus, 1758 (1)
Dasypus (Dasypus) novemcinctus Linnaeus, 1758	LC
Dasypus (Hyperoambon) beniensis Lönnberg, 1942 (2)	PE
Dasypus (Hyperoambon) kappleri Krauss, 1862 (3)	PE
Dasypus (Hyperoambon) pastasae (Thomas, 1901) (4)	PE
Dasypus (Muletia) septemcinctus Linnaeus, 1758 (5)	LC
Family Chlamyphoridae Bonaparte, 1850 (6)
Cabassous McMurtrie, 1831 (7)
Cabassous tatouay (Desmarest, 1804)	LC
Cabassous squamicaudis (Lund, 1845) (8)	PE
Cabassous unicinctus (Linnaeus, 1758)	PE
Euphractus Wagler, 1830
Euphractus sexcinctus (Linnaeus, 1758)	LC
Priodontes Cuvier, 1825
Priodontes maximus (Kerr, 1792)	VU	VU
Tolypeutes Illiger, 1811
Tolypeutes matacus (Desmarest, 1804)	NT
Tolypeutes tricinctus (Linnaeus, 1758) *	VU	EN
Order Pilosa Flower, 1883
Family Bradypodidae Gray, 1821
Bradypus Linnaeus, 1758
Bradypus torquatus Illiger, 1811 *	VU	VU
Bradypus tridactylus Linnaeus, 1758	LC
Bradypus variegatus Schinz, 1825	LC
Family Megalonychidae Gervais, 1855
Choloepus Illiger, 1811
Choloepus didactylus (Linnaeus, 1758)	LC
Choloepus hoffmanni Peters, 1858	LC
Family Cyclopedidae Pocock, 1924
CyclopesGray, 1821 (1)
Cyclopes didactylus (Linnaeus, 1758) (2)	PE
Cyclopes ida Thomas, 1900 (3)	PE
Cyclopes rufus Miranda, Casali, Perini, Machado & Santos, 2017 (4) *	PE
Cyclopes thomasi Miranda, Casali, Perini, Machado & Santos, 2017 (5)	PE
Cyclopes xinguensis Miranda, Casali, Perini, Machado & Santos, 2017 (6) *	PE
Family Myrmecophagidae Gray, 1825
Myrmecophaga Linnaeus, 1758
Myrmecophaga tridactyla Linnaeus, 1758	VU	VU
TamanduaGray, 1825
Tamandua tetradactyla (Linnaeus, 1758)	LC
Superorder Euarchontoglires
Order Primates Linnaeus, 1758
Family Aotidae Poche, 1908
Aotus Illiger, 1811
Aotus azarae (Humboldt, 1811)	LC
Aotus infulatus (Kuhl, 1820) *	NT
Aotus nancymaae Hershkovitz, 1983	VU
Aotus nigriceps Dollman, 1909	LC
Aotus trivirgatus (Humboldt, 1811)	LC
Aotus vociferans (Spix, 1823)	LC
Family Atelidae Gray, 1825
Alouatta Lacépède, 1799
Alouatta belzebul (Linnaeus, 1766) *	VU	VU
Alouatta caraya (Humboldt, 1812)	LC
Alouatta discolor (Spix, 1823) *	VU	VU
Alouatta guariba (Humboldt, 1812)	LC
Alouatta juara Elliot, 1910	LC
Alouatta macconnelli Elliot, 1910	LC
Alouatta nigerrima Lönnberg, 1941*	LC
Alouatta puruensis Lönnberg, 1941	LC
Alouatta seniculus (Linnaeus, 1766)	LC
Alouatta ululata Elliot, 1912 *	EN	EN
Ateles É. Geoffroy St.-Hilaire, 1806
Ateles belzebuth É. Geoffroy Saint-Hilaire, 1806	EN	VU
Ateles chamek (Humboldt, 1812)	EN	VU
Ateles marginatus (É. Geoffroy Saint-Hilaire, 1809) *	EN	EN
Ateles paniscus (Linnaeus, 1758)	VU
Brachyteles Spix, 1823
Brachyteles arachnoides (É. Geoffroy Saint-Hilaire, 1806) *	CR	EN
Brachyteles hypoxanthus (Kuhl, 1820) *	CR	CR
Lagothrix É Geoffroy Saint-Hilaire, 1812
Lagothrix cana (É Geoffroy Saint-Hilaire, 1812)	EN
Lagothrix lagothrica (Humboldt, 1812)	VU	VU
Lagothrix poeppigii Schinz, 1844	VU	VU
Family Callitrichidae Thomas, 1903
Callimico Miranda-Ribeiro, 1912
Callimico goeldii (Thomas, 1904)	VU
Callithrix Erxleben, 1777
Callithrix aurita (É. Geoffroy Saint-Hilaire, 1812) *	VU	EN
Callithrix flaviceps (Thomas, 1903) *	EN	EN
Callithrix geoffroyi (Humboldt, 1812) *	LC
Callithrix jacchus (Linnaeus, 1758) *	LC
Callithrix kuhlii Coimbra-Filho, 1985 *	NT
Callithrix penicillata (É. Geoffroy Saint-Hilaire, 1812) *	LC
Cebuella Gray, 1866
Cebuella niveiventris Lönnberg, 1940 (1)	LC
Cebuella pygmaea (Spix, 1823)	LC
Leontopithecus Lesson, 1840
Leontopithecus caissara Lorini & Persson, 1990 *	CR	EN
Leontopithecus chrysomelas (Kuhl, 1820) *	EN	EN
Leontopithecus chrysopygus (Mikan, 1823) *	EN	EN
Leontopithecus rosalia (Linnaeus, 1766) *	EN	EN
Mico Lesson, 1840
Mico acariensis (Roosmalen, Roosmalen, Mittermeier & Rylands, 2000) *	LC
Mico argentatus (Linnaeus, 1771) *	LC
Mico chrysoleucus (Wagner, 1842) *	LC
Mico emiliae (Thomas, 1920) *	LC
Mico humeralifer (É. Geoffroy Saint-Hilaire, 1812) *	DD
Mico humilis (Roosmalen, Roosmalen, Mittermeier & Fonseca, 1998) *	LC
Mico intermedius (Hershkovitz, 1977) *	LC
Mico leucippe (Thomas, 1922) *	VU
Mico marcai (Alperin, 1993) *	DD
Mico mauesi (Mittermeier, Schwartz & Ayres, 1992) *	LC
Mico melanurus (É. Geoffroy Saint-Hilaire, 1812)	LC
Mico munduruku Costa-Araújo, Farias & Hrbek, 2019 (2) *	PE
Mico nigriceps (Ferrari & Lopes, 1992) *	DD
Mico rondoni Ferrari, Sena, Schneider & Silva Júnior, 2010 *	VU	VU
Mico saterei (Silva Junior & Noronha, 1998) *	LC
Saguinus Hoffmannsegg, 1807
Saguinus inustus (Schwartz, 1951)	LC
Saguinus (Leontocebus) cruzlimai Hershkovitz, 1966 *	LC
Saguinus (Leontocebus) fuscicollis (Spix, 1823)	LC
Saguinus (Leontocebus) fuscus (Lesson, 1840)	LC
Saguinus (Leontocebus) nigricollis (Spix, 1823)	LC
Saguinus (Leontocebus) weddelli (Deville, 1849)	LC
Saguinus (Saguinus) bicolor (Spix, 1823) *	EN	CR
Saguinus (Saguinus) martinsi (Thomas, 1912) *	LC
Saguinus (Saguinus) midas (Linnaeus, 1758)	LC
Saguinus (Saguinus) niger (É Geoffroy Saint-Hilaire, 1803) *	VU	VU
Saguinus (Saguinus) ursula Hoffmannsegg, 1807 *	PE
Saguinus (Tamarinus) imperator (Goeldi, 1907)	LC
Saguinus (Tamarinus) labiatus (É Geoffroy Saint-Hilaire, 1812)	LC
Saguinus (Tamarinus) mystax (Spix, 1823)	LC
Family Cebidae Bonaparte, 1831
Cebus Erxleben, 1777 (3)
Cebus albifrons (Humboldt, 1812)	LC
Cebus kaapori Queiroz, 1992 *	CR	CR
Cebus olivaceus Schomburgk, 1848	PE
Cebus unicolor Spix, 1823	PE
Saimiri Vogt, 1831
Saimiri boliviensis (I. Geoffroy & Blainville, 1834)	LC
Saimiri cassiquiarensis (Lesson, 1840)	LC
Saimiri collinsi Osgood, 1916 *	PE
Saimiri macrodon Elliot, 1907	LC
Saimiri sciureus (Linnaeus, 1758)	LC
Saimiri ustus (I. Geoffroy Saint-Hilaire, 1843)	NT
Saimiri vanzolinii Ayres, 1985 *	VU	VU
Sapajus Kerr, 1792 (3)
Sapajus apella (Linnaeus, 1758)	LC
Sapajus cay (Illiger, 1815)	LC	VU
Sapajus flavius (Schreber, 1774) *	CR	EN
Sapajus libidinosus (Spix, 1823) *	LC
Sapajus macrocephalus (Spix, 1823)	LC
Sapajus nigritus (Goldfuss, 1809)	NT
Sapajus robustus (Kuhl, 1820) *	EN	EN
Sapajus xanthosternos (Wied-Neuwied, 1826) *	CR	EN
Family Pitheciidae Mivart, 1865
Cacajao Lesson, 1840 (4)
Cacajao calvus (I. Geoffroy Saint-Hilaire, 1847)	VU
Cacajao melanocephalus (Humboldt, 1812)	LC
Callicebus Thomas, 1903 (5)
Callicebus barbarabrownae Hershkovitz, 1990 *	CR	CR
Callicebus coimbrai Kobayashi & Langguth, 1999 *	EN	EN
Callicebus melanochir Wied-Neuwied, 1820 *	VU	VU
Callicebus nigrifrons (Spix, 1823) *	NT
Callicebus personatus (É. Geoffroy Saint-Hilaire, 1812) *	VU	VU
Plecturocebus Byrne, Rylands, Carneiro, Lynch Alfaro, Bertuol, da Silva, Messias, Groves, Mittermeier, Farias, Hrbek, Schneider, Sampaio & Boubli, 2016 (6)
Plecturocebus baptista Lönnberg, 1939 *	LC
Plecturocebus bernhardi Roosmalen, Roosmalen & Mittermeier, 2002 *	LC
Plecturocebus brunneus (Wagner, 1842)	LC
Plecturocebus caligatus (Wagner, 1842) (7) *	LC
Plecturocebus cinerascens (Spix, 1823) *	LC
Plecturocebus cupreus (Spix, 1823)	LC
Plecturocebus donacophilus (d’Orbigny, 1836)	LC
Plecturocebus grovesi Boubli, Byrne, da Silva, Silva-Júnior, Araújo, Bertuol, Gonçalves, Melo, Rylands, Mittermeier, Silva, Dash, Canale, Alencar, Rossi, Carneiro, Sampaio, Farias, Schneider, Hrbek, 2019 (8) *	PE
Plecturocebus hoffmannsi Thomas, 1908 *	LC
Plecturocebus moloch (Hoffmannsegg, 1807) *	LC
Plecturocebus pallescens Thomas, 1907	LC
Plecturocebus parecis Gusmão, Messias, Carneiro, Schneider, Alencar, Calouro, Dalponte, Mattos, Ferrari, Buss, Azevedo, Júnior, Nash, Rylands & Barnett, 2019 (9) *	PE
Plecturocebus stephennashi Roosmalen, Roosmalen & Mittermeier, 2002 (10) *	DD
Plecturocebus vieirai (Gualda-Barros, Nascimento & Amaral, 2012) *	DD
CheracebusByrne, Rylands, Carneiro, Lynch Alfaro, Bertuol, da Silva, Messias, Groves, Mittermeier, Farias, Hrbek, Schneider, Sampaio & Boubli, 2016 (11)
Cheracebus lucifer Thomas, 1914	LC
Cheracebus lugens (Humboldt, 1811)	LC
Cheracebus purinus Thomas, 1927 *	LC
Cheracebus regulus Thomas, 1927 *	LC
Cheracebus torquatus (Hoffmannsegg, 1807) *	LC
Chiropotes Lesson, 1840
Chiropotes albinasus (I. Geoffroy & Deville, 1848) *	EN
Chiropotes chiropotes (Humboldt, 1811)	LC
Chiropotes sagulatus (Traill, 1821)	PE
Chiropotes satanas (Hoffmannsegg, 1807) *	CR	CR
Chiropotes utahicki Hershkovitz, 1985 *	EN	VU
Pithecia Desmarest, 1804
Pithecia albicans Gray, 1860 *	LC
Pithecia cazuzai Marsh, 2014 *	DD
Pithecia chrysocephala (I. Geoffroy Saint-Hilaire, 1850)	LC
Pithecia hirsuta (Spix, 1823)	DD
Pithecia inusta (Spix, 1823)	LC
Pithecia irrorata Gray, 1843 (12)	PE
Pithecia monachus (É. Geoffroy Saint-Hilaire, 1812)	LC
Pithecia pithecia (Linnaeus, 1766)	LC
Pithecia vanzolinii Hershkovitz, 1987 (13) *	DD
Order Lagomorpha Brandt, 1855
Family Leporidae Fischer, 1817
Sylvilagus Gray, 1867
Sylvilagus brasiliensis (Linnaeus, 1758)	PE
Sylvilagus tapetillus Thomas, 1913 (1) *	PE
Order Rodentia Bowdich, 1821
Family Caviidae Fischer, 1817
Cavia Pallas, 1766
Cavia aperea Erxleben, 1777	LC
Cavia fulgida Wagler, 1831 *	LC
Cavia intermedia Cherem, Olimpio & Ximenez, 1999 *	LC	CR
Cavia magna Ximenez, 1980	LC
Galea Meyen, 1833
Galea flavidens (Brandt, 1835) *	LC
Galea spixii (Wagler, 1831)	LC
Hydrochoerus Brisson, 1762
Hydrochoerus hydrochaeris (Linnaeus, 1766)	LC
Kerodon Cuvier, 1823
Kerodon acrobata Moojen, Locks & Langguth, 1997 *	DD	VU
Kerodon rupestris (Wied-Neuwied, 1820) *	LC	VU
Family Cricetidae Fischer, 1817
Abrawayaomys Cunha & Cruz, 1979
Abrawayaomys ruschii Cunha & Cruz, 1979 *	LC
Akodon Meyen, 1833
Akodon azarae (Fischer, 1829)	LC
Akodon cursor (Winge, 1887) *	LC
Akodon lindberghi Hershkovitz, 1990 *	DD
Akodon montensis Thomas, 1913	LC
Akodon mystax Hershkovitz, 1998 *	DD	VU
Akodon paranaensis Christoff, Fagundes, Sbalqueiro, Mattevi & Yonenaga-Yassuda, 2000	LC
Akodon reigi González, Langguth & Oliveira, 1998	LC
Akodon sanctipaulensis Hershkovitz, 1990 *	DD
Akodon toba Thomas, 1921	LC
Bibimys Massoia, 1979
Bibimys labiosus (Winge, 1887)	LC
Blarinomys Thomas, 1896
Blarinomys breviceps (Winge, 1887)	LC
Brucepattersonius Hershkovitz, 1998
Brucepattersonius griserufescens Hershkovitz, 1998 *	DD
Brucepattersonius igniventris Hershkovitz, 1998 *	DD
Brucepattersonius iheringi (Thomas, 1896)	LC
Brucepattersonius nebulosus Abreu-Júnior, Vilela, Christoff, Valiati & Percequillo, 2019 (1) *	PE
Brucepattersonius soricinus Hershkovitz, 1998 *	DD
Calassomys Pardiñas, Lessa, Teta, Salazar-Bravo & Camara, 2014
Calassomys apicalis Pardiñas, Lessa, Teta, Salazar-Bravo & Camara, 2014 *	PE
Calomys Waterhouse, 1837
Calomys callidus (Thomas, 1916)	LC
Calomys callosus (Rengger, 1830)	LC
Calomys cerqueirai Bonvicino, Oliveira & Gentile, 2010 *	PE
Calomys expulsus (Lund, 1840) *	LC
Calomys laucha (Fischer, 1814)	LC
Calomys mattevii Gurgel-Filho, Feijó & Langguth 2015 (2) *	PE
Calomys tener (Winge, 1887)	LC
Calomys tocantinsi Bonvicino, Lima & Almeida, 2003 *	LC
Castoria Pardiñas, Geise, Ventura & Lessa, 2016 (3)
Castoria angustidens (Winge, 1887) (3)	PE
Cerradomys Weksler, Percequillo & Voss, 2006
Cerradomys akroai Bovincino, Casado & Weksler, 2014 *	PE
Cerradomys goytaca Tavares, Pessôa & Gonçalves, 2011 *	PE	EN
Cerradomys langguthi Percequillo, Hingst-Zaher & Bonvicino, 2008 *	PE
Cerradomys maracajuensis (Langguth & Bonvicino, 2002)	LC
Cerradomys marinhus (Bonvicino, 2003) *	LC
Cerradomys scotti (Langguth & Bonvicino, 2002)	LC
Cerradomys subflavus (Wagner, 1842)	LC
Cerradomys vivoi Percequillo, Hingst-Zaher & Bonvicino, 2008*	PE
Delomys Thomas, 1917
Delomys altimontanus Gonçalves & Oliveira, 2014 *	PE
Delomys dorsalis (Hensel, 1873)	LC
Delomys sublineatus (Thomas, 1903) *	LC
Deltamys Thomas, 1917
Deltamys araucaria Quintela, Bertuol, González, Cordeiro-Estrela, Freitas & Gonçalves, 2017 (4) *	PE
Deltamys kempi Thomas, 1917	LC
Drymoreomys Percequillo, Weksler & Costa, 2011
Drymoreomys albimaculatus Percequillo, Weksler & Costa, 2011 *	NT
Euryoryzomys Weksler, Percequillo & Voss, 2006
Euryoryzomys emmonsae (Musser, Carleton, Brothers & Gardner, 1998) *	DD
Euryoryzomys lamia (Thomas, 1901) *	VU	EN
Euryoryzomys macconnelli (Thomas, 1910)	LC
Euryoryzomys nitidus (Thomas, 1884)	LC
Euryoryzomys russatus (Wagner, 1848)	LC
Gyldenstolpia Pardiñas, D’Elía & Teta, 2009
Gyldenstolpia fronto (Winge, 1887)	CR
Gyldenstolpia planaltensis (Avila-Pires, 1972) *	PE	EN
Holochilus Brandt, 1835
Holochilus brasiliensis (Desmarest, 1819) *	LC
Holochilus chacarius Thomas, 1906	LC
Holochilus sciureus Wagner, 1842	LC
Holochilus vulpinus (Brants, 1827)	PE
Hylaeamys Weksler, Percequillo & Voss, 2006
Hylaeamys laticeps (Lund, 1840) *	NT
Hylaeamys megacephalus (Fischer, 1814)	LC
Hylaeamys oniscus (Thomas, 1904) *	NT
Hylaeamys perenensis (Allen, 1901)	LC
Hylaeamys seuanezi (Weksler, Geise & Cerqueira, 1999) (5) *	PE
Hylaeamys yunganus (Thomas, 1902)	LC
Juliomys González, 2000
Juliomys ossitenuis Costa, Pavan, Leite & Fagundes, 2007 *	PE
Juliomys pictipes (Osgood, 1933)	LC
Juliomys rimofrons Oliveira & Bonvicino, 2002 *	NT
Juliomys ximenezi Christoff, Vieira, Oliveira, Gonçalves, Valiati & Tomasi, 2016 (6) *	PE
Juscelinomys Moojen, 1965
Juscelinomys candango Moojen, 1965 *	EX	EX
Kunsia Hershkovitz, 1966
Kunsia tomentosus (Lichtenstein, 1830)	LC
Lundomys Voss & Carleton, 1993
Lundomys molitor (Winge, 1887)	LC
Microakodontomys Hershkovitz, 1993
Microakodontomys transitorius Hershkovitz, 1993 *	EN	EN
Neacomys Thomas, 1900
Neacomys amoenus Thomas, 1903 (7)	PE
Neacomys dubosti Voss, Lunde & Simmons, 2001	LC
Neacomys guianae Thomas, 1905	LC
Neacomys minutus Patton, da Silva & Malcolm, 2000 *	LC
Neacomys musseri Patton, da Silva & Malcolm, 2000	LC
Neacomys paracou Voss, Lunde & Simmons, 2001	LC
Necromys Ameghino, 1889
Necromys lasiurus (Lund, 1841)	LC
Necromys lenguarum (Thomas, 1898)	LC
Necromys urichi (Allen & Chapman, 1897)	LC
Nectomys Peters, 1861
Nectomys apicalis Peters, 1861	LC
Nectomys rattus (Pelzeln, 1883)	LC
Nectomys squamipes (Brants, 1827)	LC
Neusticomys Anthony, 1921
Neusticomys ferreirai Percequillo, Carmignotto & Silva, 2005 *	DD
Neusticomys oyapocki (Dubost & Petter, 1978)	DD
Neusticomys peruviensis (Musser & Gardner, 1974) (8)	LC
Noronhomys Carleton & Olson, 1999
Noronhomys vespuccii Carleton & Olson, 1999 *	EX
Oecomys Thomas, 1906
Oecomys auyantepui Tate, 1939	LC
Oecomys bicolor (Tomes, 1860) (9)	LC
Oecomys catherinae Thomas, 1909 (10) *	LC
Oecomys cleberi Locks, 1981 (11)	DD
Oecomys concolor (Wagner, 1845)	LC
Oecomys franciscorum Pardiñas, Teta, Salazar-Bravo, Myers & Galliari, 2016 (12)	PE
Oecomys mamorae (Thomas, 1906) (13)	LC
Oecomys paricola (Thomas, 1904) (14) *	DD
Oecomys rex Thomas, 1910	LC
Oecomys roberti (Thomas, 1904) (15)	LC
Oecomys rutilus Anthony, 1921	LC
Oecomys superans Thomas, 1911	LC
Oecomys trinitatis (Allen & Chapman, 1893)	LC
Oligoryzomys Bangs, 1900
Oligoryzomys chacoensis (Myers & Carleton, 1981)	LC
Oligoryzomys flavescens (Waterhouse, 1837)	LC
Oligoryzomys mattogrossae (Allen, 1916)	PE
Oligoryzomys messorius (Thomas, 1901)	PE
Oligoryzomys microtis (Allen, 1916)	LC
Oligoryzomys moojeni Weksler & Bonvicino, 2005 *	DD
Oligoryzomys nigripes (Olfers, 1818)	LC
Oligoryzomys rupestris Weksler & Bonvicino, 2005 *	DD	EN
Oligoryzomys stramineus Bonvicino & Weksler, 1998 *	LC
Oligoryzomys utiaritensis (Allen, 1916)	PE
Oxymycterus Waterhouse, 1837
Oxymycterus amazonicus Hershkovitz, 1994 *	LC
Oxymycterus caparaoe Hershkovitz, 1998 *	PE
Oxymycterus dasytrichus (Schinz, 1821) *	LC
Oxymycterus delator Thomas, 1903	LC
Oxymycterus inca Thomas, 1900	LC
Oxymycterus itapeby Peçanha, Quintela, Ribas, Althoff, Maestri, Gonçalves & Freitas, 2019 (16) *	PE
Oxymycterus nasutus (Waterhouse, 1837)	LC
Oxymycterus quaestor Thomas, 1903	LC
Oxymycterus rufus (G. Fischer, 1814)	LC
Phaenomys Thomas, 1917
Phaenomys ferrugineus (Thomas, 1894) *	EN
Podoxymys Anthony, 1929
Podoxymys roraimae Anthony, 1929	VU
Pseudoryzomys Hershkovitz, 1962
Pseudoryzomys simplex (Winge, 1887)	LC
Reithrodon Waterhouse, 1837
Reithrodon typicus Waterhouse, 1837	LC
Rhagomys Thomas, 1917
Rhagomys longilingua Luna & Patterson, 2003 (17)	LC
Rhagomys rufescens (Thomas, 1886) *	VU
Rhipidomys Tschudi, 1845
Rhipidomys cariri Tribe, 2005 *	DD	VU
Rhipidomys emiliae (Allen, 1916) *	LC
Rhipidomys gardneri Patton, da Silva & Malcolm, 2000	LC
Rhipidomys ipukensis Rocha, Costa & Costa, 2011 *	DD
Rhipidomys itoan Costa, Geise, Pereira & Costa, 2011 *	LC
Rhipidomys leucodactylus (Tschudi, 1845)	LC
Rhipidomys macconnelli de Winton, 1900	LC
Rhipidomys macrurus (Gervais, 1855) *	LC
Rhipidomys mastacalis (Lund, 1840) *	LC
Rhipidomys nitela Thomas, 1901	LC
Rhipidomys tribei Costa, Geise, Pereira & Costa, 2011 *	DD	EN
Rhipidomys wetzeli Gardner, 1990	LC
Scapteromys Waterhouse, 1837
Scapteromys aquaticus Thomas, 1920	LC
Scapteromys meridionalis Quintela, Gonçalves, Althoff, Sbalqueiro, Oliveira & Freitas, 2014 *	PE
Scapteromys tumidus (Waterhouse, 1837)	LC
Scolomys Anthony, 1924
Scolomys ucayalensis Pacheco, 1991	LC
Sigmodon Say & Ord, 1825
Sigmodon alstoni (Thomas, 1881)	LC
Sooretamys Weksler, Percequillo & Voss, 2006
Sooretamys angouya (Fischer, 1814)	LC
Thalpomys Thomas, 1916
Thalpomys cerradensis Hershkovitz, 1990 *	LC	VU
Thalpomys lasiotis Thomas, 1916 *	LC	EN
Thaptomys Thomas, 1916
Thaptomys nigrita (Lichtenstein, 1829)	LC
Wiedomys Hershkovitz, 1959
Wiedomys cerradensis Gonçalves, Almeida & Bonvicino, 2005 *	DD
Wiedomys pyrrhorhinos (Wied-Neuwied, 1821) *	LC
Wilfredomys Avila-Pires, 1960
Wilfredomys oenax (Thomas, 1928)	EN	EN
Zygodontomys Allen, 1897
Zygodontomys brevicauda (Allen & Chapman, 1893)	LC
Family Ctenomyidae Lesson, 1842
Ctenomys Blainville, 1826 (18)
Ctenomys bicolor Miranda-Ribeiro, 1914 *	PE	EN
Ctenomys bolivensis Waterhouse, 1848	PE
Ctenomys flamarioni Travi, 1981 *	EN	EN
Ctenomys ibicuiensis Freitas, Fernandes, Fornel & Roratto, 2012 *	PE
Ctenomys lami Freitas, 2001 *	VU	EN
Ctenomys minutus Nehring, 1887 *	DD	VU
Ctenomys nattereri Wagner, 1848 *	PE
Ctenomys rondoni Miranda-Ribeiro, 1914 *	PE
Ctenomys torquatus Lichtenstein, 1830	LC
Family Cuniculidae Miller & Gidley, 1918
Cuniculus Brisson, 1762 (19)
Cuniculus paca (Linnaeus, 1766)	LC
Family Dasyproctidae Bonaparte, 1838
Dasyprocta Illiger, 1811 (20)
Dasyprocta azarae Lichtenstein, 1823	DD
Dasyprocta croconota Wagler, 1831 *	DD
Dasyprocta fuliginosa Wagler, 1832	LC
Dasyprocta iacki Feijó & Langguth, 2013 *	DD
Dasyprocta leporina (Linnaeus, 1758)	LC
Dasyprocta prymnolopha Wagler, 1831 *	LC
Dasyprocta variegata Tschudi, 1845	DD
Myoprocta Thomas, 1803
Myoprocta acouchy (Erxleben, 1777)	LC
Myoprocta pratti Pocock, 1913	LC
Family Dinomyidae Alston, 1876
Dinomys Peters, 1873
Dinomys branickii Peters, 1873	LC
Family Echimyidae Gray, 1825
Callistomys Emmons & Vucetich, 1998
Callistomys pictus (Pictet, 1841) *	EN	EN
Carterodon Waterhouse, 1848
Carterodon sulcidens (Lund, 1838) *	DD
Clyomys Thomas, 1916
Clyomys laticeps (Thomas, 1909)	LC
Dactylomys I. Geoffroy Saint-Hilaire, 1838
Dactylomys boliviensis Anthony, 1920	LC
Dactylomys dactylinus (Desmarest, 1817)	LC
Echimys Cuvier, 1809
Echimys chrysurus (Zimmermann, 1780)	LC
Echimys vieirai Iack-Ximenes, Vivo & Percequillo, 2005 *	DD
Euryzygomatomys Goeldi, 1901
Euryzygomatomys spinosus (Fischer, 1814)	LC
Isothrix Wagner, 1845
Isothrix bistriata Wagner, 1845	LC
Isothrix negrensis Thomas, 1920 *	LC
Isothrix pagurus Wagner, 1845 *	LC
Kannabateomys Jentink, 1891
Kannabateomys amblyonyx (Wagner, 1845)	LC
Lonchothrix Thomas, 1920
Lonchothrix emiliae Thomas, 1920 *	LC
Makalata Husson, 1978
Makalata didelphoides (Desmarest, 1817)	LC
Makalata macrura (Wagner, 1842)	LC
Makalata obscura (Wagner, 1840) *	DD
Mesomys Wagner, 1845
Mesomys hispidus (Desmarest, 1817)	LC
Mesomys occultus Patton, da Silva & Malcolm, 2000 *	LC
Mesomys stimulax Thomas, 1911 *	LC
Myocastor Kerr, 1792
Myocastor coypus (Molina, 1782)	LC
Phyllomys Lund, 1839
Phyllomys blainvilii (Jourdan, 1837) *	LC
Phyllomys brasiliensis Lund, 1840 *	EN	EN
Phyllomys centralis Machado, Loss, Paz, Vieira, Rodrigues & Marinho-Filho, 2018 (21) *	PE
Phyllomys dasythrix Hensel, 1872 *	LC
Phyllomys kerri (Moojen, 1950) *	DD
Phyllomys lamarum (Thomas, 1916) *	DD
Phyllomys lundi Leite, 2003 *	EN	EN
Phyllomys mantiqueirensis Leite, 2003 *	CR
Phyllomys medius (Thomas, 1909) *	LC
Phyllomys nigrispinus (Wagner, 1842) *	LC
Phyllomys pattoni Emmons, Leite, Kock & Costa, 2002 *	LC
Phyllomys sulinus Leite, Christoff & Fagundes, 2008 *	DD
Phyllomys thomasi (Ihering, 1871) *	EN	EN
Phyllomys unicolor (Wagner, 1842) *	CR	CR
Proechimys Allen, 1899
Proechimys brevicauda (Günther, 1876)	LC
Proechimys cuvieri Petter, 1978	LC
Proechimys echinothrix da Silva, 1998 *	LC
Proechimys gardneri da Silva, 1998 *	DD
Proechimys goeldii Thomas, 1905 *	LC
Proechimys guyannensis (É. Geoffroy Saint-Hilaire, 1803)	LC
Proechimys hoplomyoides (Tate, 1939)	DD
Proechimys kulinae da Silva, 1998	DD
Proechimys longicaudatus (Rengger, 1830)	LC
Proechimys pattoni da Silva, 1998	LC
Proechimys quadruplicatus Hershkovitz, 1948	LC
Proechimys roberti Thomas, 1901 *	LC
Proechimys simonsi Thomas, 1900	LC
Proechimys steerei Goldman, 1911	LC
Thrichomys Trouessart, 1880
Thrichomys apereoides (Lund, 1839) *	LC
Thrichomys inermis (Pictet, 1843) *	LC
Thrichomys laurentius Thomas, 1904 *	DD
Thrichomys pachyurus (Wagner, 1845)	LC
Toromys Iack-Ximenes, Vivo & Percequillo, 2005
Toromys grandis (Wagner, 1845) *	LC
Trinomys Thomas, 1921 (22)
Trinomys albispinus (I. Geoffroy Saint-Hilaire, 1838) *	LC
Trinomys dimidiatus (Günther, 1877) *	LC
Trinomys elegans (Lund, 1839) *	PE
Trinomys eliasi (Pessôa & dos Reis, 1993) *	NT	VU
Trinomys gratiosus (Moojen, 1948) *	LC
Trinomys iheringi (Thomas, 1911) *	LC
Trinomys minor (dos Reis & Pessôa, 1995) *	PE
Trinomys mirapitanga Lara, Patton & Hingst-Zaher, 2002 *	DD	EN
Trinomys moojeni (Pessôa, Oliveira & dos Reis, 1992) *	EN	EN
Trinomys paratus (Moojen, 1948)	DD
Trinomys setosus (Desmarest, 1817) *	LC
Trinomys yonenagae (Rocha, 1995) *	EN	EN
Family Erethizontidae Bonaparte, 1845
Chaetomys Gray, 1843
Chaetomys subspinosus (Olfers, 1818) *	VU	VU
Coendou Lacépède, 1799
Coendou baturitensis Feijó & Langguth, 2013 (23) *	DD
Coendou bicolor (Tschudi, 1844)	LC
Coendou ichillus Voss & da Silva, 2001 (24)	DD
Coendou insidiosus (Olfers, 1818) *	LC
Coendou melanurus (Wagner, 1842)	LC
Coendou nycthemera (Olfers, 1818) *	DD
Coendou prehensilis (Linnaeus, 1758)	LC
Coendou roosmalenorum Voss & da Silva, 2001 *	DD
Coendou speratus Mendes Pontes, Gadelha, Melo de Sá, Loss, Caldara Junior, Costa & Leite, 2013 *	EN	EN
Coendou spinosus (Cuvier, 1823)	LC
Family Sciuridae Fischer, 1817
Guerlinguetus Gray, 1821
Guerlinguetus aestuans (Linnaeus, 1766)	LC
Guerlinguetus brasiliensis (Gmelin, 1788) *	PE
Hadrosciurus Allen, 1915
Hadrosciurus ignitus (Gray, 1867) (25)	LC
Hadrosciurus igniventris (Wagner, 1842)	LC
Hadrosciurus pyrrhinus (Thomas, 1898)	DD
Hadrosciurus spadiceus (Olfers, 1818)	LC
Microsciurus Allen, 1895
Microsciurus flaviventer (Gray, 1867)	LC
Sciurillus Thomas, 1914
Sciurillus pusillus (É. Geoffroy Saint-Hilaire, 1803)	LC
Superorder Laurasiatheria
Order Chiroptera Blumenbach, 1779
Family Emballonuridae Gervais, 1856
Centronycteris Gray, 1838
Centronycteris maximiliani (Fischer, 1829)	LC
Cormura Peters, 1867
Cormura brevirostris (Wagner, 1843)	LC
Cyttarops Thomas, 1913
Cyttarops alecto Thomas, 1913	LC
Diclidurus Wied-Neuwied, 1820
Diclidurus albus Wied-Neuwied, 1820	LC
Diclidurus ingens Hernández-Camacho, 1955	DD
Diclidurus isabella (Thomas, 1920)	LC
Diclidurus scutatus Peters, 1869	LC
Peropteryx Peters, 1867
Peropteryx kappleri Peters, 1867	LC
Peropteryx leucoptera Peters, 1867	LC
Peropteryx macrotis (Wagner, 1843)	LC
Peropteryx pallidoptera Lim, Engstrom, Reid, Simmons, Voss & Fleck, 2010	DD
Peropteryx trinitatis Miller, 1899	DD
Rhynchonycteris Peters, 1867
Rhynchonycteris naso (Wied-Neuwied, 1820)	LC
Saccopteryx Illiger, 1811
Saccopteryx bilineata (Temminck, 1838)	LC
Saccopteryx canescens Thomas, 1901	LC
Saccopteryx gymnura Thomas, 1901	DD
Saccopteryx leptura (Schreber, 1774)	LC
Family Furipteridae Gray, 1866
Furipterus Bonaparte, 1837
Furipterus horrens (Cuvier, 1828)	LC	VU
Family Molossidae Gervais, 1856
Cynomops Thomas, 1920
Cynomops abrasus (Temminck, 1827)	DD
Cynomops greenhalli Goodwin, 1958	LC
Cynomops milleri (Osgood, 1914)	LC
Cynomops paranus (Thomas, 1901)	DD
Cynomops planirostris (Peters, 1865)	LC
Eumops Miller, 1906
Eumops auripendulus (Shaw, 1800)	LC
Eumops bonariensis (Peters, 1874)	LC
Eumops chimaera Gregorin, Moras, Acosta, Vasconcellos, Poma, Santos & Paca, 2016 (1)	PE
Eumops delticus Thomas, 1923	DD
Eumops glaucinus (Wagner, 1843)	LC
Eumops hansae Sanborn, 1932	LC
Eumops maurus (Thomas, 1901)	DD
Eumops patagonicus Thomas, 1924	LC
Eumops perotis (Schinz, 1821)	LC
Eumops trumbulli (Thomas, 1901)	LC
Molossops Peters, 1865
Molossops neglectus Williams & Genoways, 1980	DD
Molossops temminckii (Burmeister, 1854)	LC
Molossus É. Geoffroy Saint-Hilaire, 1805
Molossus aztecus Saussure, 1860	LC
Molossus coibensis Allen, 1904	LC
Molossus currentium Thomas, 1901	LC
Molossus molossus Pallas, 1766	LC
Molossus pretiosus Miller, 1902	LC
Molossus rufus É. Geoffroy Saint-Hilaire, 1805	LC
Neoplatymops Peterson, 1965
Neoplatymops mattogrossensis (Vieira, 1942)	PE
Nyctinomops Miller, 1902
Nyctinomops aurispinosus (Peale, 1848)	LC
Nyctinomops laticaudatus (É. Geoffroy Saint-Hilaire, 1805)	LC
Nyctinomops macrotis (Gray, 1840)	LC
Promops Gervais, 1856
Promops centralis Thomas, 1915	LC
Promops nasutus (Spix, 1823)	LC
Tadarida Rafinesque, 1814
Tadarida brasiliensis (I. Geoffroy Saint-Hilaire, 1824)	LC
Family Mormoopidae Saussure, 1860
Pteronotus Gray, 1838
Pteronotus (Pteronotus) gymnonotus (Wagner, 1843)	LC
Pteronotus (Phyllodia) alitonus Pavan, Bobrowiec & Percequillo, 2018 (2)	PE
Pteronotus (Phyllodia) rubiginosus (Wagner, 1843)	LC
Pteronotus personatus (Wagner, 1843) (3)	LC
Family Natalidae Gray, 1866
Natalus Gray, 1838
Natalus macrourus (Gervais, 1856)	NT	VU
Family Noctilionidae Gray, 1821
Noctilio Linnaeus, 1766
Noctilio albiventris Desmarest, 1818	LC
Noctilio leporinus (Linnaeus, 1758)	LC
Family Phyllostomidae Gray, 1825
Ametrida Gray, 1847
Ametrida centurio Gray, 1847	LC
Anoura Gray, 1838
Anoura caudifer (É. Geoffroy Saint-Hilaire, 1818)	LC
Anoura geoffroyi Gray, 1838	LC
Artibeus Leach, 1821
Artibeus (Artibeus) fimbriatus Gray, 1838	LC
Artibeus (Artibeus) lituratus (Olfers, 1818)	LC
Artibeus (Artibeus) obscurus (Schinz, 1821)	LC
Artibeus (Artibeus) planirostris (Spix, 1823)	LC
Artibeus (Dermanura) anderseni Osgood, 1916	LC
Artibeus (Dermanura) bogotensis Andersen, 1906	LC
Artibeus (Dermanura) cinereus (Gervais, 1856)	LC
Artibeus (Dermanura) gnomus Handley, 1987	LC
Artibeus (Koopmania) concolor Peters, 1865	LC
Carollia Gray, 1838
Carollia benkeithi Solari & Baker, 2006	LC
Carollia brevicauda (Schinz, 1821)	LC
Carollia perspicillata (Linnaeus, 1758)	LC
Chiroderma Peters, 1860 (4,5)
Chiroderma doriae Thomas, 1891	LC
Chiroderma trinitatum Goodwin, 1958	LC
Chiroderma villosum Peters, 1860	LC
Choeroniscus Thomas, 1928
Choeroniscus minor (Peters, 1868)	LC
Chrotopterus Peters, 1865
Chrotopterus auritus (Peters, 1856)	LC
Desmodus Wied-Neuwied, 1826
Desmodus rotundus (É. Geoffroy Saint-Hilaire, 1810)	LC
Diaemus Miller, 1906
Diaemus youngi (Jentink, 1893)	LC
Diphylla Spix, 1823
Diphylla ecaudata Spix, 1823	LC
Dryadonycteris Nogueira, Lima, Peracchi & Simmons, 2012
Dryadonycteris capixaba Nogueira, Lima, Peracchi & Simmons, 2012	DD
Gardnerycteris Hurtado & Pacheco, 2014
Gardnerycteris crenulatum (É. Geoffroy Saint-Hilaire, 1803) (6)	LC
Glossophaga É. Geoffroy Saint-Hilaire, 1818
Glossophaga commissarisi Gardner, 1962	LC
Glossophaga longirostris Miller, 1898	LC
Glossophaga soricina (Pallas, 1766)	LC
Glyphonycteris Thomas, 1896
Glyphonycteris behnii (Peters, 1865)	DD	VU
Glyphonycteris daviesi (Hill, 1964)	LC
Glyphonycteris sylvestris Thomas, 1896	LC
Hsunycteris Parlos, Timm, Swier, Zeballos & Baker, 2014
Hsunycteris pattoni (Woodman & Timm, 2006) (7)	DD
Hsunycteris thomasi (Allen, 1904)	LC
Lampronycteris Sanborn, 1949
Lampronycteris brachyotis (Dobson, 1879)	LC
Lichonycteris Thomas, 1895
Lichonycteris degener Miller, 1931	LC
Lionycteris Thomas, 1913
Lionycteris spurrelli Thomas, 1913	LC
Lonchophylla Thomas, 1903
Lonchophylla bokermanni Sazima, Vizotto & Taddei, 1978 *	EN
Lonchophylla dekeyseri Taddei, Vizotto & Sazima, 1983 *	EN	EN
Lonchophylla mordax Thomas, 1903	NT
Lonchophylla inexpectata Moratelli and Dias (2015) (8) *	PE
Lonchophylla peracchii Dias, Esbérard & Moratelli, 2013 *	LC
Lonchorhina Tomes, 1863
Lonchorhina aurita Tomes, 1863	LC	VU
Lonchorhina inusitata Handley & Ochoa, 1997	DD
Lophostoma d’Orbigny, 1836
Lophostoma brasiliense Peters, 1866	LC
Lophostoma carrikeri (Allen, 1910)	LC
Lophostoma schulzi (Genoways & Williams, 1980)	LC
Lophostoma silvicola d’Orbigny, 1836	LC
Macrophyllum Gray, 1838
Macrophyllum macrophyllum (Schinz, 1821)	LC
Mesophylla Thomas, 1901
Mesophylla macconnelli Thomas, 1901	LC
Micronycteris Gray, 1866 (9)
Micronycteris (Xenoctenes) hirsuta (Peters, 1869)	LC
Micronycteris (Micronycteris) megalotis (Gray, 1842)	LC
Micronycteris (Micronycteris) microtis Miller, 1898	LC
Micronycteris (Schizonycteris) homezorum Pirlot, 1967	PE
Micronycteris (Schizonycteris) minuta (Gervais, 1856)	LC
Micronycteris (Schizonycteris) sanborni Simmons, 1996 *	LC
Micronycteris (Schizonycteris) schmidtorum Sanborn, 1935	LC
Mimon Gray, 1847
Mimon bennettii (Gray, 1838)	LC
Neonycteris Sanborn, 1949
Neonycteris pusilla (Sanborn, 1949)	DD
Phylloderma Peters, 1865
Phylloderma stenops Peters, 1865	LC
Phyllostomus Lacépède, 1799
Phyllostomus discolor (Wagner, 1843)	LC
Phyllostomus elongatus (É. Geoffroy Saint-Hilaire, 1810)	LC
Phyllostomus hastatus (Pallas, 1767)	LC
Phyllostomus latifolius (Thomas, 1901)	LC
Platyrrhinus Saussure, 1860
Platyrrhinus angustirostris Velazco, Gardner & Patterson, 2010	LC
Platyrrhinus aurarius (Handley & Ferris, 1972)	LC
Platyrrhinus brachycephalus (Rouk & Carter, 1972)	LC
Platyrrhinus fusciventris Velazco, Gardner & Patterson, 2010	LC
Platyrrhinus incarum (Thomas, 1912)	LC
Platyrrhinus infuscus (Peters, 1880)	LC
Platyrrhinus lineatus (É. Geoffroy Saint-Hilaire, 1810)	LC
Platyrrhinus recifinus (Thomas, 1901) *	LC
Pygoderma Peters, 1843
Pygoderma bilabiatum (Wagner, 1843)	LC
Rhinophylla Peters, 1865
Rhinophylla fischerae Carter, 1966	LC
Rhinophylla pumilio Peters, 1865	LC
Scleronycteris Thomas, 1912
Scleronycteris ega Thomas, 1912	DD
Sphaeronycteris Peters, 1882
Sphaeronycteris toxophyllum Peters, 1882	LC
SturniraGray, 1842
Sturnira (Sturnira) lilium (É. Geoffroy Saint-Hilaire, 1810)	LC
Sturnira giannae Velazco & Patterson, 2019 (10)	PE
Sturnira (Sturnira) magna De la Torre, 1966	LC
Sturnira (Sturnira) tildae De la Torre, 1959	LC
Tonatia Gray, 1827
Tonatia bidens (Spix, 1823)	DD
Tonatia maresi Williams, Willig & Reid, 1995 (11)	PE
Trachops Gray, 1847
Trachops cirrhosus (Spix, 1823)	LC
Trinycteris Sanborn, 1949
Trinycteris nicefori (Sanborn, 1949)	LC
Uroderma Peters, 1865
Uroderma bilobatum Peters, 1866	LC
Uroderma magnirostrum Davis, 1968	LC
Vampyressa Thomas, 1900
Vampyressa pusilla (Wagner, 1843)	DD
Vampyressa thyone (Thomas, 1909)	LC
Vampyriscus Thomas, 1900
Vampyriscus bidens (Dobson, 1878)	LC
Vampyriscus brocki (Peterson, 1968)	LC
Vampyrodes Thomas, 1900
Vampyrodes caraccioli (Thomas, 1889)	LC
Vampyrum Rafinesque, 1815
Vampyrum spectrum (Linnaeus, 1758)	NT
Xeronycteris Gregorin & Ditchfield, 2005
Xeronycteris vieirai Gregorin & Ditchfield, 2005 *	DD	VU
Family Thyropteridae Miller, 1907
Thyroptera Spix, 1823
Thyroptera devivoi Gregorin, Gonçalves, Lim & Engstrom, 2006	DD
Thyroptera discifera (Lichtenstein & Peters, 1855)	LC
Thyroptera lavali Pine, 1993	DD
Thyroptera tricolor Spix, 1823	LC
Thyroptera wynneae Velazco, Gregorin, Voss & Simmons, 2014	DD
Family Vespertilionidae Gray, 1821
Eptesicus Rafinesque, 1820
Eptesicus andinus Allen, 1914	LC
Eptesicus brasiliensis (Desmarest, 1819)	LC
Eptesicus chiriquinus Thomas, 1920	LC
Eptesicus diminutus Osgood, 1915	LC
Eptesicus furinalis (d’Orbigny & Gervais, 1847)	LC
Eptesicus taddeii Miranda, Bernardi & Passos, 2006 *	DD	VU
Histiotus Gervais, 1856
Histiotus alienus Thomas, 1916	DD
Histiotus diaphanopterus Feijó, Rocha & Althoff, 2015 (12)	PE
Histiotus laephotis Thomas, 1916	PE
Histiotus montanus (Philippi & Landbeck, 1861)	LC
Histiotus velatus (I. Geoffroy Saint-Hilaire, 1824)	DD
Lasiurus Gray, 1831
Lasiurus (Dasypterus) ega (Gervais, 1856)	LC
Lasiurus (Lasiurus) blossevillii (Lesson, 1826)	LC
Lasiurus (Lasiurus) castaneus Handley, 1960	DD
Lasiurus (Lasiurus) cinereus (Palisot de Beauvois, 1796)	LC
Lasiurus (Lasiurus) ebenus Fazzolari-Corrêa, 1994 *	DD
Lasiurus (Lasiurus) egregius (Peters, 1870)	DD
Lasiurus (Lasiurus) salinae Thomas, 1902	DD
Myotis Kaup, 1829
Myotis albescens (É. Geoffroy Saint-Hilaire, 1806)	LC
Myotis dinellii Thomas, 1902	LC
Myotis izecksohni Moratelli, Peracchi, Dias & Oliveira, 2011	DD
Myotis lavali Moratelli, Peracchi, Dias & Oliveira, 2011	LC
Myotis levis (I. Geoffroy Saint-Hilaire, 1824)	LC
Myotis nigricans (Schinz, 1821)	LC
Myotis riparius Handley, 1960	LC
Myotis ruber (É. Geoffroy Saint-Hilaire, 1806)	NT
Myotis simus Thomas, 1901	DD
Rhogeessa Allen, 1866
Rhogeessa hussoni Genoways & Baker, 1996	DD
Rhogeessa io Thomas, 1903	LC
Order Carnivora Bowdich, 1821
Family Canidae Fischer von Waldheim, 1817
Atelocynus Cabrera, 1940
Atelocynus microtis (Sclater, 1883)	NT	VU
Cerdocyon Smith, 1839
Cerdocyon thous (Linnaeus, 1766)	LC
Chrysocyon Smith, 1839
Chrysocyon brachyurus (Illiger, 1815)	NT	VU
Lycalopex Burmeister, 1854
Lycalopex gymnocercus (Fischer, 1814)	LC
Lycalopex vetulus (Lund, 1842) *	LC	VU
Speothos Lund, 1839
Speothos venaticus (Lund, 1842)	NT	VU
Family Mephitidae Bonaparte, 1845
Conepatus Gray, 1837
Conepatus chinga (Molina, 1782)	LC
Conepatus amazonicus (Lichtenstein, 1838) (1) *	PE
Family Mustelidae Fischer von Waldheim, 1817
Eira Smith, 1842
Eira barbara (Linnaeus, 1758)	LC
Galictis Bell, 1826
Galictis cuja (Molina, 1782)	LC
Galictis vittata (Schreber, 1776)	LC
Lontra Gray, 1843
Lontra longicaudis (Olfers, 1818)	NT
Mustela Linnaeus, 1758
Mustela africana Desmarest, 1818	LC
Pteronura Gray, 1837
Pteronura brasiliensis (Gmelin, 1788)	EN	VU
Family Otariidae Gray, 1825
Arctocephalus (G. Saint-Hilaire and F. Cuvier, 1826)
Arctocephalus australis (Zimmermann, 1783)	LC
Arctocephalus gazella (Peters, 1875)	LC
Arctocephalus tropicalis (Gray, 1872)	LC
Otaria Péron, 1816
Otaria flavescens (Shaw, 1800)	LC
Family Phocidae Gray, 1821
Hydrurga Gistel, 1848
Hydrurga leptonyx (Blainville, 1820)	LC
LobodonGray, 1844
Lobodon carcinophaga (Hombron & Jacquinot, 1842)	LC
Mirounga Gray, 1827
Mirounga leonina (Linnaeus, 1758)	LC
Family Procyonidae Gray, 1825
Bassaricyon Allen, 1876
Bassaricyon alleni Thomas, 1880	LC
Nasua Storr, 1780
Nasua nasua (Linnaeus, 1766)	LC
Potos É. Geoffroy Saint-Hilaire & Cuvier, 1795
Potos flavus (Schreber, 1774)	LC
Procyon Storr, 1780
Procyon cancrivorus (Cuvier, 1798)	LC
Family Felidae Fischer von Waldheim, 1817
Herpailurus Severtzov, 1858 (2)
Herpailurus yagouaroundi (É. Geoffroy Saint-Hilaire, 1803)	LC	VU
Leopardus Gray, 1842
Leopardus braccatus (Cope, 1889) (3)	PE
Leopardus emiliae (Thomas, 1914) (4) *	PE
Leopardus geoffroyi (d’Orbigny & Gervais, 1844)	LC	VU
Leopardus guttulus (Hensel, 1872)	LC	VU
Leopardus munoai (Ximénez, 1961) (5)	PE
Leopardus pardalis (Linnaeus, 1758)	LC
Leopardus tigrinus (Schreber, 1775)	VU	EN
Leopardus wiedii (Schinz, 1821)	NT	VU
Panthera Oken, 1816
Panthera onca (Linnaeus, 1758)	NT	VU
Puma Jardine, 1834
Puma concolor (Linnaeus, 1771)	LC	VU
Order Perissodactyla Owen, 1848
Family Tapiriidae Gray, 1821
Tapirus Brisson, 1762
Tapirus kabomani Cozzuol, Clozato, Holanda, Rodrigues, Nienow, de Thoisy, Redondo & Santos, 2013 (1)	PE
Tapirus terrestris (Linnaeus, 1758)	VU	VU
Order Artiodactyla Owen, 1848 (1)
Family Cervidae Goldfuss, 1820
Blastocerus Illiger, 1815
Blastocerus dichotomus (Illiger, 1815)	VU	VU
Mazama Rafinesque, 1817 (2)
Mazama americana (Erxleben, 1777)	DD
Mazama bororo Duarte, 1996 *	VU	VU
Mazama gouazoubira (Fischer, 1814)	LC
Mazama nana (Hensel, 1872)	VU	VU
Mazama nemorivaga (Cuvier, 1817)	LC
Odocoileus Rafinesque, 1832
Odocoileus virginianus (Zimmermann, 1780)	LC
Ozotoceros Ameghino, 1891
Ozotoceros bezoarticus (Linnaeus, 1758)	NT	VU
Family Tayassuidae Palmer, 1897
Pecari Reichenbach, 1835
Pecari tajacu (Linnaeus, 1758)	LC
Tayassu Fischer, 1814
Tayassu pecari (Link, 1795)	VU	VU
Family Balaenidae Gray, 1821
Eubalaena Gray, 1864
Eubalaena australis (Desmoulins, 1822)	LC	EN
Family Balaenopteridae Gray, 1864
Balaenoptera Lacépède, 1804
Balaenoptera acutorostrata Lacépède, 1804	LC
Balaenoptera bonaerensis Burmeister, 1867	NT
Balaenoptera borealis Lesson, 1828	EN	EN
Balaenoptera edeni Anderson, 1879	LC
Balaenoptera musculus (Linnaeus, 1758)	EN	CR
Balaenoptera physalus (Linnaeus, 1758)	VU	EN
Megaptera Gray, 1846
Megaptera novaeangliae (Borowski, 1781)	LC
Family Delphinidae Gray, 1821
Cephalorhynchus Gray, 1846
Cephalorhynchus commersonii (Lacépède, 1804)	LC
Delphinus Linnaeus, 1758
Delphinus delphis Linnaeus, 1758	LC
Feresa Gray, 1870
Feresa attenuata Gray, 1874	LC
Globicephala Lesson, 1828
Globicephala macrorhynchus Gray, 1846	LC
Globicephala melas (Traill, 1809)	LC
GrampusGray, 1828
Grampus griseus (Cuvier, 1812)	LC
Lagenodelphis Fraser, 1956
Lagenodelphis hosei Fraser, 1956	LC
Lagenorhynchus Gray, 1846
Lagenorhynchus australis (Peale, 1848)	LC
Lissodelphis Gloger, 1841
Lissodelphis peronii (Lacépède, 1804)	LC
Orcinus Fitzinger, 1860
Orcinus orca (Linnaeus, 1758)	DD
Peponocephala Nishiwaki & Norris, 1966
Peponocephala electra (Owen, 1846)	LC
Pseudorca Reinhardt, 1862
Pseudorca crassidens (Owen, 1846)	NT
Sotalia Gray, 1866
Sotalia fluviatilis (Gervais & Deville, 1853)	DD
Sotalia guianensis (Van Bénéden, 1864)	NT	VU
Stenella Gray, 1866
Stenella attenuata (Gray, 1846)	LC
Stenella clymene (Gray, 1850)	LC
Stenella coeruleoalba (Meyen, 1833)	LC
Stenella frontalis (Cuvier, 1829)	LC
Stenella longirostris (Gray, 1828)	LC
Steno Gray, 1846
Steno bredanensis (Cuvier, 1828)	LC
Tursiops Gervais, 1855 (3)
Tursiops truncatus (Montagu, 1821)	LC
Family Iniidae Gray, 1846
Inia d’Orbigny, 1834
Inia araguaiaensis Hrbek, Farias, Dutra & da Silva, 2014 (4) *	PE
Inia boliviensis d’Orbigny, 1834	PE
Inia geoffrensis (Blainville, 1817)	EN	EN
Family Kogiidae Gill, 1871
Kogia Gray, 1846
Kogia breviceps (Blainville, 1838)	DD
Kogia sima (Owen, 1866)	DD
Family Phocoenidae Gray, 1825
Phocoena Cuvier, 1816
Phocoena dioptrica Lahille, 1912	LC
Phocoena spinipinnis Burmeister, 1865	NT
Family Physeteridae Gray, 1821
Physeter Linnaeus, 1758
Physeter macrocephalus Linnaeus, 1758	VU	VU
Family Pontoporiidae Gray, 1870
Pontoporia Gray, 1846
Pontoporia blainvillei (Gervais & d’Orbigny, 1844)	VU	CR
Family Ziphiidae Gray, 1865
Berardius Duvernoy, 1851
Berardius arnuxii Duvernoy, 1851	DD
Hyperoodon Lacépède, 1804
Hyperoodon planifrons Flower, 1882	LC
Mesoplodon Gervais, 1850
Mesoplodon densirostris (Blainville, 1817)	DD
Mesoplodon europaeus (Gervais, 1855)	DD
Mesoplodon grayi von Haast, 1876	DD
Mesoplodon hectori (Gray, 1871)	DD
Mesoplodon layardii (Gray, 1865)	DD
Mesoplodon mirus True, 1913	DD
Ziphius Cuvier, 1823
Ziphius cavirostris Cuvier, 1823	LC

Publication Dates

• Publication in this collection
  17 Aug 2020
• Date of issue
  2020

History

• Received
  27 Aug 2019
• Accepted
  2 Dec 2019