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The Vomeronasal Organ of New World Monkeys (Platyrrhini).

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THE ANATOMICAL RECORD 294:2158–2178 (2011)
The Vomeronasal Organ of New World
Monkeys (Platyrrhini)
School of Physical Therapy, Slippery Rock University, Slippery Rock, Pennsylvania
Department of Anthropology, University of Pittsburgh, Pittsburgh, Pennsylvania
Department of Anthropology, The Graduate Center at the City University of New York,
New York
New York Consortium in Evolutionary Primatology (NYCEP), New York, New York
Department of Anatomical Sciences and Neurobiology, University of Louisville School of
Medicine, Louisville, Kentucky
Dallas World Aquarium, Dallas, Texas
Department of Biology, Slippery Rock University, Slippery Rock, Pennsylvania
Department of Anatomy, Physiology, and Pharmacology, College of Veterinary Medicine,
Auburn University, Auburn, Alabama
Central Oregon Community College, 2600 N.W. College Way, Bend, Oregon
Although all platyrrhine primates possess a vomeronasal organ
(VNO), few species have been studied in detail. Here, we revisit the
microanatomy of the VNO and related features in serially sectioned samples from 41 platyrrhine cadavers (14 species) of mixed age. Procedures
to identify terminally differentiated vomeronasal sensory neurons (VSNs)
via immunolabeling of olfactory marker protein (OMP) were used on
selected specimens. The VNO varies from an elongated epithelial tube
(e.g., Ateles fusciceps) to a dorsoventrally expanded sac (e.g., Saguinus
spp.). The cartilage that surrounds the VNO is J-shaped or U-shaped in
most species, and articulates with a groove on the bony palate. Preliminary results indicate a significant correlation between the length of this
groove and length of the VNO neuroepithelium, indicating this feature
may serve as a skeletal correlate. The VNO neuroepithelium could be
identified in all adult primates except Alouatta, in which poor preservation prevented determination. The VNO of Ateles, described in detail for
the first time, had several rows of VSNs and nerves in the surrounding
lamina propria. Patterns of OMP-reactivity in the VNO of perinatal platyrrhines indicate that few or no terminally differentiated VSNs are present at birth, thus supporting the hypothesis that some platyrrhines may
have delayed maturation of the VNO. From a functional perspective, all
platyrrhines studied possess structures required for chemoreception
(VSNs, vomeronasal nerves). However, some microanatomical findings,
such as limited reactivity to OMP in some species, indicate that some lineages of New World monkeys may have a reduced or vestigial vomeronasal
C 2011 Wiley Periodicals, Inc.
system. Anat Rec, 294:2158–2178, 2011. V
Grant sponsor: National Science Foundation; Grant number:
BCS 0820751; Grant sponsor: Department of Homeland
Security; Grant number: 01-G-022.
*Correspondence to: Timothy D. Smith, School of Physical
Therapy, Slippery Rock University, Slippery Rock, PA. Tel: 724738-2885. E-mail:
Received 15 September 2011; Accepted 16 September 2011
DOI 10.1002/ar.21509
Published online 1 November 2011 in Wiley Online Library
Key words: olfaction;
vomeronasal neuroepithelium; VNO; platyrrhines
The vomeronasal organ (VNO) is the peripheral receptor organ for the vomeronasal, or accessory, olfactory
system. Experimental work has demonstrated that this
chemosensory system is an important mediator of sociosexual behaviors in many mammals. Disrupting the
vomeronasal system (VNS), through removal of the VNO
or lesions of vomeronasal nerves, reduces some sexual
and aggressive behaviors in many rodents (Powers and
Winans, 1975; Beauchamp et al., 1982; Clancy et al,
1984; Lepri and Wysocki, 1987; Wysocki et al., 1991),
opossums (Jackson and Harder, 1996), and at least one
primate (Aujard, 1997). Yet, it has become clear that the
VNO is not a critical mediator of these behaviors in all
mammals (Barrett et al., 1993; Woodley et al., 2004).
One fundamental difference between Old World monkeys and New World monkeys is the complete absence of
the VNO in the former (Frets, 1912). The VNO, comprising a neuroepithelial tube in most mammals, disappears
during fetal development in Old World monkeys (Hendrickx, 1971; Wilson and Hendrickx, 1977). Although a
vestigial VNO has been described in some other catarrhines (e.g., humans and chimpanzees—Smith et al.,
2001b, 2002; Witt et al., 2002), not even a trace of the
VNO neuroepithelium has been found in postnatal Old
World monkeys (Jordan, 1972; Smith et al., 2001a,b). In
contrast, the VNO of at least some New World monkeys
(platyrrhines) possesses bipolar neurons and vomeronasal nerves, characteristics in keeping with a functional neuroepithelium (Taniguchi et al., 1992; Mendoza
et al., 1994; Dennis et al., 2004).
Regarding differences in olfactory anatomy, Frets
(1912) observed that in monkeys ‘‘ : : : nature has made
an experiment’’ (p. 137). For Frets, this experiment could
potentially reveal the function of the VNS, since it is
present in one group and absent in the other. Nearly a
century later, the function of the VNO continues to be
debated (Halpern and Martı́nez-Marcos, 2003; Doty,
2010). Among primates, the platyrrhine–catarrhine dichotomy has generated debate concerning evolution of
olfactory abilities (Liman and Innan, 2003; Zhang and
Webb, 2003; Gilad et al., 2004; Wang et al., 2010; Matsui
et al., 2010; Young et al., 2010), yet has yielded little
insight into the function of the VNO itself. Attempts to
experimentally ablate the VNO have produced no
evidence, as of yet, for VNS function in platyrrhines
(Barrett et al., 1993).
The present state of knowledge on the VNS of primates, then, is incomplete. Aside from quantitative studies
of the accessory olfactory bulb (Stephan et al., 1981), the
site of first order vomeronasal sensory neuron synapses,
the VNS has been studied in only a few platyrrhine species. Anatomical knowledge of the VNO, the peripheral
receptor organ for this neural system, is especially lacking in platyrrhines. Only small-bodied species have provided the basis for detailed microscopic descriptions
(Taniguchi et al., 1992; Mendoza et al., 1994; Dennis
et al., 2004).
Herein, we present new findings on the VNO and
related structures in 14 species of platyrrhines. The
aims of this report are two-fold. First, this is the broadest taxonomic survey of VNO morphology in platyrrhines
to date. Very few species have been described regarding
neuroepithelial organization. The present report
includes observations on the vomeronasal neuroepithelium (VNNE) morphology in 14 species, including three
for which VNNE has never been described using adult
specimens. New and existing information is then summarized in an effort to reconcile previous descriptions of
the VNO in platyrrhines, which have included vague
descriptors such as ‘‘well developed’’ (Hershkovitz, 1977;
Bhatnagar and Meisami, 1998; Brennan, 2001; Cartmill
and Smith, 2009), or as showing ‘‘signs of reduction’’
(Maier, 1980; p. 229). A second aim is to assess maturational characteristics of the VNNE in perinatal monkeys.
Specifically, the present study seeks to investigate the
organization of the VNNE and histochemistry and immunoreactivity of vomeronasal sensory neurons (VSNs)
and extraepithelial neuronal bodies surrounding the
VNO. The hypothesis that platyrrhine primates exhibit
delayed maturation of the VNO (Evans and Grigorieva,
1994; Shimp et al., 2003) is tested using the perinatal
Sample Characteristics
The present study is based on observations on 41
adult, juvenile, perinatal, and fetal specimens from 14
species of primates (Table 1). The sample includes some
specimens that were used previously (Smith et al., 2002,
2003a, 2004; Dennis et al., 2004). Included are three species that have never been subjected to detailed examination using adult samples, Ateles fusciceps, Cebuella
pygmaea, and Leontopithecus rosalia. Tables 2–4 present
a compilation of new and previously published observations (e.g., VNO measurements, immunoreactivity to
neuronal markers, or conventional histochemical procedures), and includes an update on metric and non-metric
aspects of the VNO in platyrrhini.
All specimens were acquired from cadaveric remains
preserved in 10% formalin. This opportunistic sample
included animals that were stillborn or died postnatally
in captivity at various research centers and zoos (Table
1). No animal was euthanized specifically for the present
study. Whenever possible, both males and females of
each genus were studied.
Sample Preparation
Procedures for sectioning material used in earlier
studies are described elsewhere (see Smith et al., 2003a
and Dennis et al., 2004 for details). Newly prepared histological specimens were processed similarly, described
briefly as follows. Before embedding, cranial length
TABLE 1. Species, sample size, and source of primates sectioned
Species (common name)
Aotus trivirgatus (Northern owl monkey)
Alouatta caraya (black howler monkey)
Alouatta seniculus (red howler monkey)
Ateles fusciceps (brown-headed spider monkey)
Callithrix jacchus (common marmoset)
Cebuella pygmaea (3) (pygmy marmoset)
Leontopithecus rosalia (golden lion tamarin)
Pithecia pithecia (2) (White-faced saki)
Saimiri boliviensiss (Bolivean squirrel monkey)
Saguinus spp
Sample size, age (sex)
2 adults (m, f)
1 adult (m)
1 fetus (f)
1 adult (f)
2 adults; 6 perinatal (m, f)
3 adults (3f); 1 41-day-old (f); 1 perinatal (f)
2 adults (m,f); 1 juvenile; 1 perinatal
1 adult (m), 1 perinatal (u)
1 adult (f); 6 perinatal (m, u)
6 adults (m,f); 2 juveniles; 2 perinatal (m,f)
CMZ, Cleveland Metroparks Zoo; DWA, Dallas World Aquarium; GPZ, Gladys Porter Zoo; MKCMR, Michael E Keeling
Center for Comparative Medicine and Research ; NEPC; New England Primate Center; NMNH, National Museum of Natural History; PZ, Philadelphia Zoo; USAPRC, University of South Alabama Primate Research Center; WRPC, Wisconsin Regional Primate Center.
TABLE 2. Summary of characteristics of the vomeronasal complex in Platyrrhines: Duct
communications and cartilaginous capsule
Aotus trivirgatus
Alouatta caraya
Ateles fusciceps
Callithrix jacchus
Cebuella pygmaea
Cebus spp
Leontopithecus spp
Pithecia pithecia
Saimiri spp.
Saguinus spp.
VNO duct communica.
dorsoventrally elongate
dorsoventrally elongate?
round or dorsoventrally
round or dorsoventrally
irregular/ dorsoventr. elongate
irregular/ dorsoventr. elongate
dorsal NPD
dorsal NPD
dorsal NPD
mid NPD
mid NPD
mid NPD
mid to dorsal NPD
dorsal NPD
mid NPD
mid NPD
‘‘closed’’ J
‘‘open’’ J compressed
closed J
closed J
open J compressed
closed J
open J
open J
closed J
1, 2
1, 3
4, 5
1, 3
2, 3
NPD, nasopalatine duct; VNC, vomeronasal cartilage; VNO, vomeronasal organ; References: 1, this study; 2, Hunter et al.,
1984; 3, Smith et al., 2003a; 4, Frets, 1912; 5, Jordan, 1968
TABLE 3. Summary of characteristics of the vomeronasal complex in Platyrrhines: Lamina propria
Aotus trivirgatus
Alouatta caraya
Ateles fusciceps
Callithrix jacchus
Cebuella pygmaea
Cebus spp
Leontopithecus spp
Pithecia pithecia
Saimiri spp.
Saguinus spp.
Neural elements
Glands, commun.
Glands, histochem.
large and small branches
smaller branches
large and small branches, PVNG
large and small branches
‘‘nerve bundle’’
small branches
small branches
small branches, PVNG
A, P, D,V
A, P, R,V, L
A, P, D, V, M, L
ciliated ducts
A, P, D, M, L
ciliated ducts
1, 2, 4
PVNG, paravomeronasal ganglia; Glands, commun. (refers to VNO surface at which glands communicate, and other comments): A, anterior; P, posterior: D, dorsal; V, ventral; M, medial; L, lateral. References: 1, this study; 2, Smith et al., 2002;
3, Frets (1912) describes and shows a drawing of Cebus with a large nerve bundle near the VNO; 4, Smith et al., 2003a.
(prosthion-inion) and palatal length (prosthion-posterior
mid-palatal point) were measured with digital calipers
to the nearest 0.01 mm. In some specimens, damage during necropsy prevented acquisition of these measurements. All paraffin embedded heads were serially
sectioned at 10–12 lm and stained for histomorphometric analysis using ImageJ software (NIH). At least every
tenth section was mounted on glass slides with serial
numbers and stained with hematoxylin–eosin and
Gomori trichrome procedures. Intervening sections were
saved for alternative procedures.
Histochemical and Immunohistochemical
Procedures for the Study of Subadults
A subset of the perinatal sample (Leontopithecus,
Saguinus) was studied to elucidate characteristics of cell
bodies associated with nerves adjacent to the VNO. Since
similar cells were previously demonstrated to react to the
lectin Ulex europaeus-1 (UEA-1) (Evans and Grigorieva,
1994) as well as neuron-specific markers (Dennis et al.,
2004; Smith et al., 2004), we selected closely adjacent
unstained sections of tamarin VNOs to characterize
TABLE 4. Measurements and microanatomical characteristics of the vomeronasal neuroepithelium in
adult Platyrrhines
VNNEL (mm)
VNEA (mm2)
Lumen/VNO ratio
Aotus trivirgatus
Alouatta caraya
Ateles fusciceps
Callithrix jacchus
Cebuella pygmaea
Leontopithecus rosalia
Pithecia pithecia
Saimiri spp.
Saguinus spp.
2.6, 2.8
2.7, 3.1
0.04, 0.066
0.038, 0.04
0.40, 0.49
0.45, 0.49
P (LM)
P (LM)
P (LM)
1, 2
1, 2
3, 4
1, 5
1, 2
1, 2, 3, 4
VNNEL, anteroposterior length of VNNE; VNEA, cross-sectional area of VNNE (in mm2) ¼ an average of the cross-sectional area at 25th, 50th, and 75th percentile of VNNE length; SEO, sensory epithelium only; ISE, interrupted sensory epithelium; ?, unclear from available specimens; P, present; LM, light microscopic observation; TEM, transmission electron
microscopic observations.
() indicates that no OMP-reactive cells are visible in the VNNE (only one specimen of Saimiri boliviensis was studied by
Smith et al., 2011); (þ), indicates sparse, individual OMP-reactive cells visible in the VNNE (see Smith et al., 2011 for
details); (þþ), indicates multiple rows of OMP-reactive cells visible in the VNNE (see Smith et al., 2011 for details).
References: 1, This study; 2, Smith et al., 2011; 3, Taniguchi et al., 1992; 4, Smith et al., 2004; 5, unpublished observations (on Cebuella only).
groups of cells. In addition, we examined VNNE immunoreactivity to olfactory marker protein (OMP), a marker of
terminally differentiated neurons. These specimens were
examined to see whether a broad range of platyrrhines
resemble Saguinus spp. (Dennis et al., 2004; Smith et al.,
2011) in having few or no mature VSNs at birth.
Lectin binding was performed using the avidin-biotin
peroxidase complex (ABC) method following specified
procedures. Protocols were adopted and modified accordingly from Vector literature and previous studies
(Nakajima et al., 1998). All procedures were done at
room temperature and solutions were mixed daily just
prior to and/or during experimental procedures.
After sections were deparaffinized and hydrated in
graded ethanols, they were rinsed in running tap water
for 5 min, incubated in 1% bovine serum albumin (BSA)
for 30 min, rinsed in 0.05 M tris buffered saline (TBS, pH
¼ 7.2) for 5 min, and incubated in UEA-1 (Vector Inc,
Burlingame, CA) for 30 min. Following incubation, sections were washed in 0.05 M TBS for 5 min, incubated
with ABC reagent for 30 min, washed in 0.05 M TBS for
5 min, reacted with diaminobenzidine (DAB) labeling kit
(Vector Inc.) for 5 min, and washed in distilled water for
15 min. Sections were dehydrated and cleared with xylene in the reverse order of that noted above and
mounted with Permount (Fisher Scientific, Pittsburgh,
PA). Several slides were lightly counterstained with
hemotoxylin prior to this step for delineation and confirmation of adjacent cell populations.
Two control conditions were examined to confirm lectin specificity: (1) omission of lectin which was replaced
by 0.05 M TBS for 30 min; and, (2) introduction of 0.4 M
of inhibitory sugar (L-Fucose, from Ferro Pfanstiehl,
Waukegan, IL), premixed with lectin, for 30 min.
For neuron-specific beta tubulin (BT) (Covance
Inc.,Princeton, NJ) immunohistochemistry, sections were
deparaffinized in Hemo-D (Scientific Safety Products,
Fort Lauderdale, FL) and hydrated to distilled H2O
(dH2O). Endogenous peroxidase-like activity was blocked
by holding the slides for 20 min in 0.9% H2O2 made in
absolute methanol. After washing in dH2O, the slides
were washed 3 3 min each in phosphate buffered saline (PBS, pH 7.4). Subsequently, the slides were placed
in humidified chambers and incubated 30 min in blocker
solution of 5% normal horse serum and 2.5% BSA. The
slides were then rinsed briefly in PBS and returned to
their chambers. Anti-BT (Covance Inc.), diluted 1:16,000
in blocker, was applied to the tissues and the slides were
left overnight at 23*C. Next morning, the slides were
washed 3 3 min each in PBS, and the tissues were
incubated 1 hr in biotinylated horse anti-mouse IgGs,
(Vector, Inc.), diluted 1:200 in blocker. Following incubation in the secondary antibody, the tissues were washed
2 3 min each in PBS and incubated 30 min in ABC reagent (Vector Inc.). After 2 3 min washes in PBS, antibody binding was detected with a DAB labeling kit.
Secondary antibody binding specificity was tested by
running tissue sections of each species through all the
steps above except that the sections were incubated in
blocker without primary antibody.
Sections selected for double labeling with BT and PGP
9.5 (Chemicon, Temecula, CA) were prepared as above
except that the incubation in methanol and H2O2 was
omitted. Anti-BT (1:2,000) and anti-PGP (1:1,000) antibodies were applied as a cocktail and treated as
described above. Secondary antibodies conjugated to
Alexa fluorochromes (Molecular Probes, Carlsbad, CA)
were also applied as a cocktail. The slides were mounted
with Vectashield (Vector, Inc.) and viewed with a Nikon
E600 microscope equipped with epifluoresence optics.
Images were made with a Spot Slider digital camera and
Spot Advanced (4.0.1) software (Diagnostic Instruments,
Sterling Heights, MI).
For OMP immunohistochemistry, sections were deparaffinized in Hemo-D (Scientific Safety Products, Fort
Lauderdale, FL) and hydrated to distilled H2O (dH2O).
Endogenous peroxidase-like activity was blocked by
holding the slides for 20 min in 0.9% H2O2 made in
absolute methanol. After washing the slides in dH2O,
the slides were washed 3 3 min each in PBS (pH 7.4).
Subsequently, the slides were placed in humidified
chambers and incubated overnight in anti-OMP (Wako
Chemicals USA, Inc, Richmond, VA) diluted appropriately in blocker (5% normal rabbit serum and 2.5% BSA
in PBS). The next morning, slides were washed 3 3
min each in PBS, and the tissues were incubated 1 hr in
biotinylated rabbit anti-goat IgG (Vector Inc.), diluted
1:200 in blocker. Following incubation in the secondary
antibody, the tissues were washed 2 3 min each in PBS
and incubated 30 min in ABC reagent. After 2 3 min
washes in PBS, antibody binding was detected with a
DAB labeling kit. Secondary antibody binding specificity
was tested by running tissue sections of each species
through all the steps above, except that the sections
were incubated in blocker without primary antibody.
Slides were examined using light microscopy with a
Leica DMLB photomicroscope at 100 to 630. VNOs
were studied for reactivity in apical processes, presumptive receptor (bipolar) cells, supporting cells, and basal
The Vomeronasal Groove
Posterior to its communication to the nasopalatine
duct (Fig. 1A), the VNO lies in a cartilaginous capsule
that articulates with the maxillary bone (Fig. 1B). A
groove forming from the articulation of the vomeronasal
cartilage and the hard palate provides a good approximation of VNO gross dimensions such as length and
width, based on studies of primates including strepsirrhines and platyrrhines (Garrett et al., 2009; Garrett,
2010). In the present study, we examined the relationship between the vomeronasal groove (VNG) and the
VNO in platyrrhines only. In particular, VNG length and
width were regressed against a measurement of the sensory part of the VNO, length of the VNNE.
Linear measurements were collected on photographs
of previously sectioned platyrrhine nasal fossae using
ImageJ software. For length of the VNG, the number of
slides in which the VNNE and VNG were observed in
contact were counted and multiplied by slice thickness.
Width measurements were taken of the VNG (Fig. 1B)
at the 25th, 50th, and 75th percentiles of the length of
the VNO using the line tool in ImageJ.
This regression analysis was necessarily preliminary,
since our sample was limited to specimens with a VNNE
that was identifiable across anteroposterior limits. Fifteen adult specimens including nine species were suitable for the analysis.
General Observations on Adult Sample
In all adult specimens, except where freezing artifact
prevented assessment, three cell types are identifiable in
the VNNE. Although ultrastructural confirmation of
cells types was not made, these matched previous
descriptions of supporting, basal, and receptor cells in
the VNNE. A row of presumptive supporting cells with
abundant apical cytoplasm is seen adjacent to the
lumen. Presumptive basal cells are visible as small, flat,
or triangular cells with scant cytoplasm that are
sparsely distributed along the basement membrane. Presumptive VSNs are visible in all adult specimens (except
in Alouatta caraya and Pithecia pithecia due to freezing
damage). In previous studies, the affinity of cells within
the VNNE to neuron-specific markers was described in
Fig. 1. Vomeronasal complex of Cebuella pygmaea. A: View of the
ductal communication (*) of the VNO to the nasopalatine duct (npd).
B: Location of the VNO within a scroll of cartilage (vnc) that articulates
with the maxillary bone (m). The cartilage rests in a groove (width indicated by yellow arrows) in the maxilla. C: View of the VNO, showing a
prominent large bundle of vomeronasal nerves (double arrows). Note
the proportionally small VNO lumen (L). D: Higher magnification view
of the VNO neuroepithelium (vnne), showing the multiple rows of VSNs
(arrows). Gomori trichrome. gd, gland duct; nf, nasal fossa. Scale
bars: A, 400 lm; B, 1 mm; C, 200 lm; D, 30 lm.
some adult callitrichines (Dennis et al., 2004; Smith
et al., 2004), verifying that a neuronal complement of
cells exists in VNNE of at least some species.
Specific Observations on Adult Sample, by
Cebuella. The VNO of Cebuella pygmaea communicates with the nasopalatine duct at a mid-palatal level
(Fig. 1A). The VNO itself is round or oval in cross-section,
with large bundles of vomeronasal nerves connected dorsally (Fig. 1B,C). The VNNE has rows of three or more
VSN nuclei on all sides of the VNO in cross-section (Fig.
1C). Glands communicate to the VNO lumen, from anterior, posterior, medial, lateral, and dorsal sides of the
VNO (Fig. 1C,D). Preliminary observations using OMP
immunohistochemistry indicate the VNNE of adult
Cebuella is OMP (), whereas olfactory neuroepithelium
Fig. 2. Vomeronasal complex of Leontopithecus rosalia. A–D: Anteroposterior views of the VNO, its ductal communication (*) to the
nasopalatine duct (npd), its cartilaginous capsule (vnc), and communicating glands (gl). E: The dorsoventrally elongate VNO, with a large
lumen (L). Numerous venous sinuses surround it. F: The VNO neuroepithelium (vnne) is visible on both sides of the organ, although there
appear to be gaps (brackets) where receptor neurons are lacking. G–
F: Higher magnification views showing that one to two rows of VSNs
(arrows). Small bundles of nerves (double arrows) connect to the base
of the VNNE (2G, I). Microvilli (open arrows, 2H) are visible. Basal cell
(bc) and supporting cell (sc) nuclei are also apparent. A: Hematoxylin–
eosin; B–I: Gomori trichrome. Scale bars: A–D, 400 lm; E, 200 lm; F,
50 lm; G, 30 lm; H, 20 lm; I, 20 lm.
of the same specimens is OMP (þ) (unpublished
times irregular and always dorsoventrally elongated sac
(Fig. 2C–E). Anteriorly, the irregular border has fingerlike projections in all directions, where glands communicate from all sides (Fig. 2C). Posterior to the VNO, more
glands are found (Fig. 2D), connecting to the VNO via
multiple ducts.
Numerous venous sinuses surround the VNO (Fig.
2E). The walls of the VNO are neuroepithelial on both
sides (Fig. 2F), although only one to two rows of receptor
nuclei are visible (Fig. 2G–I). VSNs are sparsely
Leontopithecus. The VNO, lined with neuroepithelium, extends anterior to its communication with the
nasopalatine duct (Fig. 2A, also showing anterior glands
on the contralateral side). The VNO opens into the nasopalatine duct via a short, laterally projecting duct (Fig.
2B). Posterior to this, the VNO becomes a large, some-
Fig. 3. Vomeronasal complex of Aotus trivirgatus. A–D: Anteroposterior views of the VNO, its ductal communication (*) to the nasopalatine
duct (npd), its cartilaginous capsule (vnc), and communicating glands
(gl). E: The dorsoventrally elongate VNO, with a large lumen (L). Numerous venous sinuses (vs) surround it. F: The VNO neuroepithelium (vnne)
is visible on both sides of the organ. E: Shows gland ducts (gd) entering
the VNO dorsally and ventrally. F: Higher magnification view showing
rows of VSNs (arrows). Small bundles of nerves (double arrows) connect to the base of the VNNE. A–D: Gomori trichrome; E,F: Hematoxylin
eosin. Scale bars: A–D, 500 lm; E, 125 lm; F, 30 lm.
distributed in some regions, with some isolated patches
having no apparent VSNs at all (Fig. 2F,H). Long microvilli are visible at the apical surface of the VNNE (Fig.
2H). Vomeronasal nerves are visible in the lamina propria (Fig. 2G,I).
VNNE (Fig. 3F). Nuclei of VSNs typically are arranged
in rows of three.
Aotus. In Aotus, glands located anterior to the VNO
and in the region of its communication to the nasopalatine duct (Fig. 3A) open into the VNO. The VNO is dorsoventrally elongate in its cross-sectional contour (Fig.
3B,C) and ends posteriorly in communication with
glands (Fig. 3D). Throughout its length, numerous
glands communicate at dorsal and ventral poles (Fig.
3E). Posteriorly, the vomeronasal cartilage is partially
ossified, especially at posterior levels (Fig. 3C,D).
The VNNE is present at all surfaces in the coronal
plane, and nerve bundles are visible in contact with the
Saimiri. The VNO communicates with the nasopalatine duct at the mid-level of the palate in the single
adult specimen studied (Fig. 4A), among adjacent glands
that open into the VNO. The VNO is round in cross-section (Fig. 4B), ending in communication with posterior
clusters of glands (Fig. 4C).
In the lamina propria, numerous venous sinuses are
present (Fig. 4D) and small bundles of nerve fibers are
seen at high magnification (Fig. 4E). The VNNE has one
to three rows of VSNs (Fig. 4E). At intermediate and
posterior levels, gland ducts are ciliated (Fig. 4F,G).
Ateles. Anterior to the VNO communication with the
nasopalatine duct, the vomeronasal cartilage mostly
Fig. 4. Vomeronasal complex of Saimiri sciureus. A–C: Anteroposterior views of the VNO, its ductal communication (*) to the nasopalatine duct (npd), its cartilaginous capsule (vnc), and communicating
gland ducts (gd). D: The round VNO, with a large lumen (L). Numerous
venous sinuses (vs) surround it. Note the uniform thickness of the wall
of the VNO. E: The VNO neuroepithelium has several rows of VSNs
and also shows a gland duct (gd) entering the VNO. Very small nerves
bundles (nn), containing few axons each, are visible in the lamina
propria (E). F,G: Show ciliated (open arrows) gland ducts that entered
the VNO at approximate mid-levels as well as posteriorly. All stained
with Gomori trichrome except (F) (hematoxylin–eosin). Scale bars: A–
C, 400 lm; D, 200 lm; E, 20 lm; F, 40 lm; G, 20 lm.
Fig. 5. Vomeronasal complex of Ateles fusciceps. A–C: Anteroposterior views of the VNO, its ductal communication (*) to the nasopalatine duct (npd), its cartilaginous capsule (vnc), and communicating
gland ducts (gd). D: The VNO is dorsoventrally compressed (C) and
ends as a large duct receiving gland ducts (gd) from all sides (D,E) as
well as posteriorly. The lumen (L) is large but varies in relation to thickness of the VNO wall. F: The VNO neuroepithelium (vnne) has three or
more rows of VSNs (arrows). Small bundles of axons (nn) are visible in
the lamina propria. A,B,F: Gomori trichrome; (C–E) hematoxylin–eosin.
Scale bars: A–C, 800 lm; D, 100 lm; E, 50 lm; F, 30 lm.
encircled small glands (Fig. 5A), which communicated by
ducts to the VNO lumen. A stratified cuboidal duct
extended for a short distance from the upper (nasal)
extent of the nasopalatine duct to the VNO itself (Fig.
5B), here defined as beginning where a neuroepithelium
was present. The VNO is dorsoventrally compressed
through most of its extent (Fig. 5C), with glands entering posteriorly, superiorly, inferiorly, medially, and laterally through the wall of the VNO (e.g, Fig. 5D,E). In the
single adult Ateles specimen, the VNNE appears to have
distinct junctions with non-sensory epithelium (Fig.
6B,C). Long microvilli are visible (Fig. 6C). Some surfaces of the VNNE are covered with a fine poorly defined
meshwork (Fig. 6A). These may represent apical cell surface projections, but could not be identified at 1000 levels of light microscopy.
ing artifact) in these specimens was noted, but only the
surrounding lamina propria was described in detail. In
A. caraya, the VNO is dorsoventrally elongate in crosssection and communicates with the NPD at about the
mid-palate (Fig. 7). Compared to all other specimens,
this specimen’s vomeronasal cartilage is atypical because
it remains superior to the nasal septum’s base and has a
discontinuous bar-shaped form at some cross-sectional
intervals (Fig. 7B–D). Glands communicate with the
VNO at multiple sites (Fig. 7A–E). In P. pithecia (not figured), the VNO is round in cross-section and the opening
of the VNO occurs in the nasopalatine duct, but very
close to the base of the nasal cavity.
Observations on Subadult Samples
Alouatta and Pithecia. The general morphology of
General observations on perinatal and fetal
samples. Perinatal or fetal platyrrhine primates have
adult Alouatta caraya and Pithecia pithecia were previously described. In the work by Smith et al. (2002), the
poor preservation of the VNO epithelium (due to freez-
in common a relatively thin VNNE (Fig. 8A,C,E,G). The
VNNE of perinatal marmosets (e.g., Callithrix, Fig. 8B),
has no discernable differentiation of VSN and supporting
cell nuclei. The VNNE is only one to two nuclei in depth.
In Saimiri boliviensis and Pithecia pithecia, most of the
VNNE is two to three nuclear rows in depth, but a row
of cells with abundant apical cytoplasm (presumptive
supporting cells) borders the lumen (Fig. 8D,F). Some
patches of thicker epithelium are sparsely distributed in
each of these species and these patches contain VSN-like
cell clusters (Fig. 8D,F). Preservation of the fetal
Alouatta seniculus was poorer than the perinatal specimens. However, existing tissues indicate a similarity to
Saimiri and Pithecia, and it is possible that a more discrete VNNE exists in this specimen (Fig. 8G,H).
Lectin and immunohistochemical results on
subadult primates. No OMP reactivity is observed in
the VNO of perinatal monkeys, including marmosets
(Callithrix jacchus and Cebuella pygmaea) (Fig. 9) or the
Saki monkey (Pithecia pithecia, not shown). The olfactory neuroepithelium of these perinatal monkeys is OMP
(þ) (Fig. 9). In one perinatal squirrel monkey, no OMP
(þ) VSNs are visible. In a second Saimiri specimen,
which is older (10 days postnatal), a single layer of OMP
(þ) cells is seen in the VNNE. The olfactory neuroepithelium of this specimen is also OMP (þ), with rows of reactive olfactory sensory neurons (Fig. 10).
In subadult Saguinus geoffroyi (Fig. 11) and Leontopithecus rosalia (not shown), nearly the entire VNO epithelium is intensely UEA-1 (þ) (Fig. 11C,E), but BT (þ)
cells in adjacent sections are sparse (Fig. 11B,G). In L.
rosalia, more numerous BT (þ) cells are seen (Fig.
12b,e) compared to S. geoffroyi. Double labeling with BT/
PGP 9.5 revealed that neonatal S. geoffroyi had mostly
BT()/ PGP 9.5(þ) cells in the VNO epithelium (Fig.
11D) while in L. rosalia colabeling was seen at perinatal
and juvenile ages (Fig. 12c,f).
Extraepithelial PGP 9.5 (þ) or BT(þ) cell bodies are
found in association with the VNO or nerves to the VNO
in both species of tamarin at perinatal and older subadult ages (Figs. 11D,G and 12b,e). The lectin procedure
reveals that cell bodies found in vomeronasal nerves are
either UEA-1() or weakly UEA-1(þ). In both species of
tamarin, some small UEA-1(þ) cell bodies are found
within nerves connected to the VNO (e.g., Fig. 11E).
Double labeling with BT/PGP 9.5 in L. rosalia reveals
that populations of cells clustered within the VNN are
BT(þ)/PGP 9.5(þ) (Fig. 12c,f).
Control sections in which UEA-1 was preabsorbed
with 0.4 M of L-fucose showed minimal to no reactivity.
Additionally, controls where no lectin was introduced
(0.05 M TBS replaced lectin) demonstrated no VNO
staining specificity.
Vomeronasal Groove
Fig. 6. Higher magnification views of the VNNE in Ateles fusciceps.
A: Rows of up to four nuclei of VSNs (arrows) are shown. Also note a
layer of stained structures at the surface of the VNNE, which could be
short microvilli (double arrows). B: A transition from VNNE to a thinner
epithelium is shown. C: At some locations fine, long microvilli (long
arrows) were visible at the apical surface of the VNO epithelium. A,B:
Gomori trichrome; (C) hematoxylin–eosin. bc, basal cell nucleus; sc,
supporting cell nucleus. Scale bars: 20 lm.
Dimensions of the VNG are significantly correlated
with VNNE length, despite the small sample size (Fig.
13). VNG width is significantly correlated with VNNE
length (r2 ¼ 0.613, P < 0.05). The correlation is stronger
if Ateles fusciceps is removed from the analysis (r2 ¼
0.798, P < 0.01). The location of Ateles in the plot distribution indicates this species falls below the regression
line for the remaining, smaller species (Fig. 13). Further
analysis may determine whether the relationship of
VNG width to VNNE width differs in larger relative to
smaller platyrrhines. There is a stronger association of
Fig. 7. Vomeronasal complex of Alouatta caraya. A–E: An adult A.
caraya. A–E: Anteroposterior views of the VNO, its ductal communication (*) to the nasopalatine duct (npd), its cartilaginous capsule (vnc),
and communicating gland ducts (gd) are shown. Poor preservation
prevented a description of epithelial structure, but a dorsoventrally tall
contour is apparent in this specimen (E). Also note the diminutive
VNC. All stained with Gomori trichrome except C (hematoxylin–eosin).
Scale bars: A–D, 500 lm; E, 250 lm.
Fig. 8. Vomeronasal complex of perinatal and fetal platyrrhines.
The bottom row shows an enlargement of boxed regions in the top
row. Perinatal Callithrix jacchus (A,B), Saimiri sciureus (C,D), and Pithecia pithecia (E,F) are shown. G,H: Fetal specimen of Alouatta seniculus. In all species, VSNs (arrows) were sparse, or in discontinuous
clusters. B: Shows a cluster of cells dorsal to the VNO that may represent a paravomeronasal ganglia. A,B,G,H: Gomori trichrome; (C–F) hematoxylin–eosin. gd, gland duct; L, lumen of VNO; vncC, vomeronasal
cartilage; vno, vomeronasal organ. Scale bars: A, C, E 500 lm; G, 200
lm; B, D, F, H, 30 lm.
VNG length with VNNE length (r2 ¼ 0.772, P < 0.01;
Fig. 13).
across the order, rather than a single evolutionary trajectory for all primates (Smith and Rossie, 2006; Smith
et al., 2007). Taken together, anatomical and genetic
data provide one coherent evolutionary picture, supporting a hypothesis that the catarrhine VNS became nonfunctional prior to the divergence of the cercopithecoid
and hominoid lineages (Smith et al., 2001a; Liman and
Innan, 2003; Zhang and Webb, 2003; Rossie, 2005; Bhatnagar and Smith, 2006). For the main olfactory system
(MOS), different lines of evidence support remarkably
different evolutionary views on primate olfaction (Cave,
The two olfactory systems (main olfactory system and
VNS) are generally considered reduced in primates compared to other mammals (Stephan et al., 1981; Barton,
2006), yet recent findings on the VNO and other olfactory structures indicate a complex pattern of reduction
Fig. 9. Expression of OMP in perinatal callitrichines. A: VNO of Callithrix, and (B) Cebuella, showing lack of OMP expression in the wall
of the VNO. C,D: OMP expression in the olfactory neuroepithelium
(OE) of Callithrix (same specimen as Fig. 11A) showing reactivity of ol-
factory receptor neurons (arrows) and olfactory nerves (ON). E: OMP
expression in the olfactory neuroepithelium (OE) of Cebuella (same
specimen as Fig. 11B) showing reactivity of olfactory neuroepithelium
(OE) and olfactory bulb (OB). Scale bars: A–D, 20 lm; E, 250 lm.
1973; Rouquier et al., 2000; Whinnett and Mundy, 2003;
Laska et al., 2005, 2006). For example, data on relative
volume of the main olfactory bulb indicate that primates
may vary in the extent that they rely on the MOS and
VNS for dietary versus social functions (Alport, 2004).
Anatomical and genetic evidence have generated several evolutionary scenarios for the primate VNS (Fig.
14) that hinge entirely on extant taxa. Smith et al.
(2007) suggested that details of evolutionary change
may be gleaned from anatomic variations of olfactory
structures in extant primates, especially since these var-
iations may bear on interpretations of fossil primate
morphology and behavior. However, the VNO of platyrrhines, a major radiation and one that is generally more
primitive than cercopithecoids and hominoids in numerous features, has not been described in similar terms in
all previous investigations.
Maier (1980) provided a functional assessment of the
VNS in platyrrhines primarily based on Saimiri and callitrichines (species not specified). He described a basic
similarity of the VNO epithelium to olfactory neuroepithelium (i.e., the presence of supporting, receptor, and
Fig. 10. Expression of OMP in a 10-day-old Saimiri boliviensis. A: VNO, and (B) olfactory neuroepithelium. A single row of OMP(þ) cells is seen along most of the VNO wall [arrows, (A)]. Multiple rows
of OMP(þ) cells are seen in the olfactory neuroepithelium [arrows, (B)] on, olfactory nerves. Scale bars:
100 lm.
basal cells). Subsequently, this arrangement was confirmed in Aotus trivirgatus (Hunter et al., 1984), Saguinus fuscicollis (Hunter et al., 1984; Mendoza et al.,
1994), and Callithrix jacchus (Taniguchi et al., 1992;
Dennis et al., 2004). Transmission electron microscopy
specifically confirmed that receptor neurons are present
in two species of callitrichines (Taniguchi et al., 1992;
Mendoza et al., 1994). Based on the presence of a neuroepithelium, some authors concluded that the VNO of certain platyrrhines may be functional (Maier, 1980;
Taniguchi et al., 1992).
However, potential signs of evolutionary regression
in the platyrrhine VNS have also been observed. In
comparison to Tupaia and strepsirrhines, Maier (1980)
discussed ‘‘signs of reduction’’ in the VNO of Saimiri
and, to a greater degree, in unspecified callitrichines.
In this regard, he noted the lack of large venous
sinuses, the lack of a distinction between non-sensory
and sensory portions of the VNO, and a poor distinction of cell types within some portions of the VNO epithelium. Smith et al. (2003a) noted that the VNO of
Saguinus geoffroyi comprises a neuroepithelium interrupted by patches of nonsensory epithelium; sparse
distribution of VSNs was confirmed by immunohistochemical studies on this species (Dennis et al., 2004;
Smith et al., 2004). Hunter et al. described the VNO of
Ateles geoffroyi as ‘‘similar to that lining the nasopalatine ducts’’ (p. 223, 1984) without making reference to
sensory cells. Thus, it remains uncertain whether all
platyrrhines possess a neuroepithelial VNO and to
what extent it varies among taxa. Below, we integrate
data from our present report with existing literature
on the primate VNNE.
General Characteristics of the Vomeronasal
Complex in Platyrrhines
VNO communication with the oral and nasal
cavities. The basic route of stimulus access to the
VNO, via the nasopalatine duct, has been established
previously (Jordan, 1972; Hunter et al., 1984; Shimp
et al., 2003). In this study, we found that the specific
point of communication with the nasopalatine duct
varies. In Leontopithecus, Aotus, Ateles, Alouatta, and
especially in Pithecia, the VNO communicates with the
dorsal part of the nasopalatine duct.
A vomeronasal duct, which is a non-sensory stratified
cuboidal or stratified squamous epithelial conduit connecting the sensory part of the VNO to the nasopalatine
duct (see Shimp et al., 2003), is barely discernable as a
distinct conduit in most species. Only larger species,
such as Ateles fusciceps, have a vomeronasal duct that
was distinct from the VNNE for multiple serial sections.
In most species, a neuroepithelium is present near the
communication point of the VNO with the nasopalatine
VNO shape. The cross-sectional contour of the VNO
is round or oval in some small platyrrhines (Cebuella,
Callithrix and Saimiri), and dorsoventrally elongate in
Saguinus, Leontopithecus, and Aotus (Table 2, and references therein). In two larger platyrrhines, Cebus (Jordan, 1972) and Ateles (Table 2), the VNO is round or
dorsoventrally compressed. In one adult Alouatta caraya, the VNO is dorsoventrally elongated (although
freezing may have distorted the region).
Fig. 11. In neonatal Saguinus geoffroyi (A–C), hematoxylin and eosin stained sections [(A), inset with position indicated by open arrow]
showed an epithelium of the VNO that was nearly homogeneous,
showing patches of non-sensory epithelium (nse), and thicker, possibly
sensory epithelium (double arrows). B: BT reactivity was restricted to
single (double arrows) or sparsely scattered cells in neonates in each
section of the VNO. C: UEA-1 reactivity was ubiquitously present
throughout the VNO epithelium. No BT or UEA-1 axons were seen in
the lamina propria adjacent to the VNO. D: Shows a neonatal S. geoffroyi prepared with PGP 9.5 (green)/BT (red), showing the numerous
PGP 9.5þ cells in the VNO epithelium (double arrows) and one extrae-
pithelial reactive cell (open arrow). The VNO of 1-month-old (D–F)
Saguinus geoffroyi has ubiquitous UEA-1 reactivity throughout the
VNO epithelium, except for scattered weakly or non-reactive cells
(white arrows). The vomeronasal nerves (VNN) are non-reactive but
sometime contain UEA-1þ cells (open arrows). A more posterior section of the VNO is shown in (E) (Gomori Trichrome) in which the VNN
can be seen superiorly to the VNO. F: Shows a higher magnification
of an adjacent section, in which extraepithelial BTþ neuronal bodies of
different sizes and morphology can be seen near the VNN. L ¼ lumen.
Scale bars, A–C and inset, 100 lm; D, 20 lm (inset, 500 lm); E, G ¼
50 lm; F ¼ 100 lm.
Relative size of the lumen varies. Marmosets have the
smallest proportional size of the lumen while Aotus and
Saguinus spp. have the largest (see lumen/VNO ratios,
Table 4). Tamarins in particular have proportionally
large VNOs, in part because the lumen is voluminous.
The VNNE of tamarins is also more superoinferiorly
elongated compared to other platyrrhines. Because of
that, it has a greater cross-sectional area in the coronal
plane (Table 4). The platyrrhine VN varies from an elongated tube (Ateles) to a mediolaterally compressed sac
(Saguinus, Leontopithecus, and Aotus).
marsupials, insectivores, most bats, and many primates
(Broom, 1897; Cooper and Bhatnagar, 1976; Sánchez Villagra, 2001; Wöhrmann-Repenning and Bergmann,
2001). Here, we refer to this morphology as an open
J configuration (Figs. 1–3; Table 2). Some mammals possess more complete, laterally enclosed capsules, which
may be mostly or entirely osseous (Salazar and Sanchez
Quintero, 1998). This specialization of the vomeronasal
capsule, in which the vomeronasal cartilage is mostly
replaced by extensions of the vomer bone (Salazar and
Sanchez Quinteiro, 1998), may maximize efficiency of
the vasomotor vomeronasal pump mechanism, which
delivers stimuli to the lumen of the VNO (Meredith and
O’Connell, 1979). The nearly complete enclosure of the
VNO by cartilage in Saguinus (Maier, 1980; Hunter
et al., 1984) and Aotus (Hunter et al., 1984) has been
noted previously. Here we document this closed J morphology as being characterstic of Aotus and most callitrichine genera (Table 2; note Callimico has not been
VNO capsule and glandular apparatus. The
vomeronasal capsule shows two variants. In one morph,
the cartilage is open laterally, that is, there is no cartilaginous (or osseous) wall between the dorsolateral side
of the VNO and the respiratory mucosa (Fig. 4). This
morphology is fairly typical of many mammals, including
Fig. 12. The vomeronasal organ (VNO) of a neonatal (a–c) and juvenile (d–f) L. rosalia, illustrating populations of extraepithelial cell bodies
at both ages [open arrows, (a,b,e)]. Adjacent sections prepared with
BT/PGP 9.5 are shown in (c) and (f). These double-labeled fluores-
cence (BTþ ¼ red; PGP 9.5þ ¼ green; co-labeled ¼ yellow) images
indicated that such cells were BTþ/PGPþ (open arrows). L ¼ lumen;
VNC ¼ vomeronasal cartilage; VNN ¼ vomeronasal nerves. Scale
bars: a, b, 100 lm; c, e, f, 50 lm; d, 200 lm.
Fig. 13. Plots of VNG width and VNG length against VNNE Length.
Linear regression line is indicated, although the largest species (Ateles
fusciceps) indicates some scaling phenomena require further investigation. Af ¼ Ateles fusciceps, At ¼ Aotus trivirgatus, Cj ¼ Callithrix
jacchus, Cp ¼ Cebuella pygmaeus, Lr ¼ Leontopithecus rosalia, Sb ¼
Saguinus bicolor, Sg ¼ Saguinus geoffroyi, So ¼ Saguinus oedipus,
Saim ¼ Saimiri boliviensis.
Fig. 14. Cladogram showing major primate groups with detailed
relationships amongst the living platyrrhini (constructed based on information from Steiper and Ruvolo (2003), Ray et al. (2005), and
Hodgson et al., (2009)). Cladogram also traces character history for
VNO character states using parsimony analysis (Mesquite software).
Groups with bullet have known morphology. A VNO with well-defined
neuroepithelial and non-sensory parts was likely primitive for Euprimates. A VNO with sensory-epithelium only was most likely primitive
for haplorhines based on its extant distribution. The ancestral VNO
state of catarrhines is equivocal, with a SEO, displaced, or absent
VNO being equally likely.
studied). The capsule appears to ossify in Aotus, which
has also been reported for ‘‘tamarins’’ by Maier (1980,
taxa not specified).
The platyrrhine VNO receives glandular ducts at multiple points along its anteroposterior, mediolateral, and
dorsoventral axes (Table 3). All species studied here
have multiple entry points for glands at more than two
surfaces, as described previously for some individual
species (Jordan, 1972; Mendoza et al., 1984). Glands at
the dorsal poles extend out of the vomeronasal capsule
in all platyrrhines. This is common among mammals
(Roslinski et al., 2000), and may represent a primitive
mammalian innovation in which some septal glands are
co-opted into the VNS (Smith and Bhatnagar, 2009). All
species also receive gland ducts posteriorly. The anterior
communicating glands in Alouatta are not unique, as
once written by Smith et al. (2002), and may be a similarly widely distributed trait. At least one other gland
entry point (medial, lateral, and or ventral) was
observed in every species examined in this study.
In many mammals, the VNO comprises two tissues: a
sensory epithelium and a receptor-free (non-sensory) epithelium. Gland duct entry points appear to occur commonly at the junction of these two epithelia (Roslinski
et al., 2000). Since all platyrrhines possess a VNO with
multiple glandular inputs, this may well be a primitive
arrangement for a haplorhine common ancestor. This is
in keeping with the vestigial VNOs of some (possibly all)
hominoids, in which the VNO itself may serve mainly as
a glandular duct (Smith et al., 2002). This vestige, which
is ciliated (like some of the gland ducts of platyrrhines)
may well have evolved from the primitive haplorhine
VNO as a remnant duct that delivers secretions to the
proportionally large homninoid nasal fossa.
Structure of the Vomeronasal Epithelium
Based on prominent adjacent and apparently communicating vomeronasal nerves, Frets (1912) suspected the
VNO in adult Cebus was neurosensory in nature. The
results of the present study, combined with previous
findings, indicate that a neuroepithelial VNO exists in
most, if not all, platyrrhines (Table 2). Some previous
reports could not confirm the presence of neuroepithelium, but this very likely relates to artifactual phenomena. One methodological consideration that arises in
histological studies of rare samples, such as non-human
primates, is that it is difficult to obtain adequately prepared samples for study. Thus, postmortem decay or
freezing have interfered with assessments in some studies (Smith et al., 2002). In addition, Hunter et al. (1984)
suggested prolonged decalcification may have distorted
the epithelial structure in one specimen of Ateles, a dilemma that is particular to larger species. We have
achieved our best results with a formic acid–sodium citrate solution (see above) which has been effective in
decalcifying very large tissue blocks from hominoids
(Smith et al., 2002) and relatively large monkeys such
as Ateles. This decalcifier works slowly (taking more
than two months in some cases) and appears to leave
epithelial tissue undistorted.
Most evidence indicates that a more or less homogeneous epithelial wall is the primitive VNO epithelial morphology for anthropoids (Fig. 14). This is in contrast to
the ventromedially restricted VNNE seen in most mammals, including strepsirrhine primates (Hunter et al.,
1984; Smith and Bhatnagar, 2009; Smith et al., 2011). A
uniform sensory epithelium was described previously for
most New World primates (Maier, 1980; Hunter et al.,
1984; Taniguchi et al., 1992; Mendoza et al., 1994). This
morphology typifies adult tarsiers as well (Starck, 1975;
Wöhrmann-Repenning and Bergmann, 2001), raising the
Non-sensory epithelium has been described in the VNO
of some platyrrhines, but in no case have descriptions
matched the morphology of receptor-free epithelium. Receptor-free epithelium (Breipohl et al., 1979) is widespread among therians. It is often ciliated, and may
therefore play a role in transporting secretions within the
VNO. To date, two types of distribution of non-sensory epithelium have been reported in adult platyrrhines, neither of which matches the descriptions of receptor-free
epithelium by Breipohl et al. (1979). First, large patches
of non-sensory tissues have been described in Cebus (Jordan, 1972) and Ateles (this study). In Cebus capuchinus,
Jordan (1972) described some medial to lateral differences of the VNO wall. Medially, sensory epithelium was
observed. Laterally, sensory epithelium was more limited
and instead a stratified, non-sensory epithelium predominated. In Ateles fusisceps, a ventral patch of non-sensory
epithelium is described here. Based on such small samples, it is unclear whether these represent species characteristics or aberrations. Transient stages of VNO
development have been observed in which the VNO of
haplorhines has a non-sensory component (e.g., tamarins,
Dennis et al., 2004; tarsiers and squirrel monkeys, Smith
et al., 2003b). Thus, the morphology could be atavistic.
A second example of non-sensory epithelium pertains
to the so-called ‘‘interrupted sensory epithelium’’
described in Saguinus geoffroyi by Smith et al. (2003a).
In this case, the VNO wall has numerous interruptions of
the neuroepithelium by intervening patches of non-sensory epithelium. Immunohistochemical studies using
neuron-specific markers indicate that the basis for this
morphology may be a reduced density of VSNs, which
occur in clusters (Dennis et al., 2004; Smith et al., 2004).
Leontopithecus may also possess this phenotype. Of all
species examined in this study, adult Leontopithecus
exhibits the fewest rows of VSNs (one to two). A single
specimen of Leontopithecus chrysomelas similarly was
observed to have only few VSNs that react to antibodies
for neuron-specific BT, when compared to marmosets or a
strepsirrhine species (Smith et al., 2004). Taken together,
these results suggest that the VSN population is small in
Saguinus and Leontopithecus, and patches of non-sensory
epithelium may simply reflect a regional dearth of VSNs.
Based on available evidence, therefore, the occurrence
of non-sensory components does not constitute a functional receptor-free epithelium (Breiphol et al., 1979) as
observed in other mammals. If not an atavistic trait,
such non-sensory patches could be construed as a
byproduct of neural reduction. Marmoset only possesses
one type of G-protein in its accessory olfactory bulb
(Takagami et al., 2004). In rodents and marsupials, two
distinct populations of G-protein-expressing cells (Go
and Gi2) are found in the VNNE and AOB (Halpern and
Martı́nez-Marcos, 2003). Therefore, it is possible that in
marmosets an entire subset of VSNs is absent. If this
applies to platyrrhines broadly, it may explain the relatively small number of rows of VSNs that are found in
the VNNE compared to strepsirrhines (Smith et al.,
2004) and other mammals (Halpern and Martı́nez-Marcos, 2003).
Previous findings showed that some tamarins (Saguinus spp.) have a very sparse distribution of VSNs, as
detected by neuronal markers such as PGP 9.5 and neuron-specific BT (Dennis et al., 2004; Smith et al., 2004).
Since no estimates of total neuronal numbers have been
provided to date, it is not known whether Saguinus spp.
have fewer as opposed to more widely dispersed VSNs.
However, recent work has shown primate VNOs vary in
reactivity to an additional neuronal marker, OMP (Dennis et al., 2004; Smith et al., 2005, 2011). Since this
marker is expressed in terminally differentiated olfactory or VSNs (Farbman and Margolis, 1980; Weiler and
Benali, 2005), the limited reactivity to OMP in the VNO
of some platyrrhines indicates the presence of only few
mature sensory neurons (Dennis et al., 2004; Smith
et al., 2011). Smith et al. (2011) examined five species of
Saguinus, and found this genus has few mature vomeronasal neurons at any age when compared to strepsirrhines. Although other platyrrhine species have only
been investigated using one or two specimens, some
resemble Saguinus (e.g., Aotus and Saimiri) while
others appear to have a VNNE with multiple rows of
OMP(þ) VSNs (Dennis et al., 2004; Smith et al., 2011).
In possessing relatively small numbers of terminally differentiated VSNs, Saguinus spp (and possibly other platyrrhines as well) differ from a diverse array of
mammals, including some other primates (Halpern and
Martı́nez-Marcos, 2003; Smith et al., 2005, 2011). Observations on a larger sample of platyrrhines are still much
needed; available observations of atelines are extremely
limited and no detailed information exists regarding the
VNNE of adult pitheciines.
Ontogenetic Characteristics of the Vomeronasal
Delayed maturation of the VNNE. Perinatal morphology of the VNO is, to date, documented in callitrichines alone. Evans and Grigorieva (1994) suggested
tamarins (Saguinus labiatus) and marmosets (Cebuella
pygmaea) have delayed VNNE maturation. This hypothesis, based on lectin histochemistry (Evans and Grigorieva, 1994), was supported in subsequent studies using
neuron-specific markers. In Leontopithecus rosalia and
Saguinus geoffroyi, the density of VSNs detected by PGP
9.5, neuron-specific BT, or OMP is lower than in lemurs
at birth (Dennis et al., 2004; Smith et al., 2004). In our
sample, we detected no OMP reactivity in the VNOs of
marmosets and Pithecia. Our limited sample suggests
that infant Saimiri boliviensis may have mature VSNs,
as also observed in juvenile Leontopithecus rosalia. However, this is difficult to interpret without additional
specimens of other ages, especially since an adult of this
species was observed to lack OMP(þ) VSNs (Smith
et al., 2011).
For most species, the lack of perinatal expression of
OMP provides support to the hypothesis of delayed maturation of the VNNE. However, if the VNNE is poorly
developed in adults of certain species, as suggested for
Saguinus spp. and possibly other primates (Aotus and
Saimiri) (Smith et al., 2004, 2011; this study) delayed
maturation appears to be a misnomer. In such species,
the neuroepithelial part of the VNS may be postnatally
reduced or even vestigial, unless OMP is not required
for VNO function (Smith et al., 2011).
In callitrichines, in particular, the hypothesis of
delayed maturation may be applied to Callithrix, which
has a thin OMP() VNNE at birth, and later develops a
thick VNNE with terminally differentiated VSNs (Taniguchi et al., 1992; Dennis et al., 2004). Saguinus has a
different postnatal developmental trend, in which the
density of VSNs may actually decrease postnatally. In S.
geoffroyi, OMP(þ) VSNs are scarce in both newborns
and adults (Dennis et al., 2004; Smith et al., 2011). The
density of BT(þ) VSNs decreases postnatally, based on a
small sample of Saguinus spp. (Smith et al., 2004).
Although previous reports indicate OMP reactivity is
greater in a juvenile L. rosalia compared to perinatal
specimens, observations in this study suggest VSNs may
become sparsely distributed in adult L. rosalia. The possibility that this reflects a lifelong comparatively low
rate of neurogenesis in Saguinus and Leontopithecus
could be investigated with certain neuronal markers in
the future (e.g., Gap43).
Another goal of the present study was to examine late
fetal/perinatal representatives across subfamilies to complement the published data on tamarins. Though our
sample was limited for some taxa (with no Callicebus or
Aotus), a coherent pattern among all specimens was a
thin VNO epithelium with a patchy distribution of VSNlike cells. The available specimens suggested two distinct groups. First, callitrichines appear to have especially poorly differentiated VNOs at birth compared to
perinatal Pithecia, Saimiri and fetal Alouatta. In all callitrichines, there is no distinction (at least by light microscopy and the procedures used here) between
supporting cells and other types at birth; VSN-like cells
are distributed as isolated cells (as confirmed based on
the distribution of BT(þ) cells—Smith et al., 2004). P.
pithecia and S. boliviensis have groupings of VSN-like
cells scattered in clusters. Although its preservation was
not optimal, the fetal A. seniculus appeared to have
larger patches of neuroepithelium as well.
Extraepithelial neuronal bodies adjacent to
the VNNE. Evans and Grigorieva (1994) suggested
that neuron-like cells in the lamina propria surrounding
the VNO of callitrichines were evidence for prolonged
postnatal neurogenesis. These authors speculated that
some of these cells were migratory, possibly luteinizing
hormone releasing hormone (LHRH) neurons. Subsequent reports identified neuronal bodies in the lamina
propria surrounding the VNO of Saguinus spp., Leontopithecus rosalia, and Callithrix jacchus (Dennis et al.,
2004; Smith et al., 2004). Using Saguinus and Leontopithecus, we confirm here that cell bodies associated with
nerves adjacent to the VNO are both UEA (þ) and reactive to neuronal markers.
Although it is not clear that these cells are migratory
or ganglionic, their neuronal identity is certain. In morphology, these PGP 9.5 and/or BT reactive cells varied
from relatively large-bodied neurons, as described for
the paravomeronasal ganglia of bats (Bhatnagar and
Kallen, 1974, Bhatnagar et al., 2006) to relatively fusiform neurons with processes of varying length, as
described in LHRH neurons of mice (Wu et al., 1997).
Our observation of UEA-1 (þ) cell bodies within the
vomeronasal nerves also is noteworthy since such cells
have been observed in close association with LHRH neurons in other mammals (Tobet et al., 1997), and UEA-1
is thought to identify one of the neural cell adhesion
molecules (Pestean et al., 1995). Nonetheless, the presence of a migratory population of neurons would seem
extremely prolonged in callitrichines compared to other
mammals described to date. At least some neuroblastic
cell migration from olfactory epithelium occurs postnatally in rodents (Monti-Graziadei, 1992). Postnatal
migration also may occur along the vomeronasal nerves
of rodents, yet it is clear that there is a marked postnatal reduction of LHRH reactive cells found in the
nasal cavity from about mid-gestation (77.6% in nasal
region/22.4% in CNS) to birth (7.9% in nasal region/
92.1% in CNS) (Wu et al., 1997). Similarly, large numbers of extraepithelial neurons associated with the VNO
have not previously been described in primates, except
during embryonic and fetal stages (Boehm and Gasser,
1993; Kjær and Fischer Hansen, 1996; Smith et al.,
A second explanation is that these neurons represent
aberrant ganglia, as previously described for cranial
nerves (Satomi & Takahashi, 1986). OMP expression was
reported before in subepithelial nasal ganglia (Storan
and Key, 2006). In this regard, the frequent presence of
paravomeronasal ganglia in adult bats, another taxonomic group with extreme variations in the VNO (Bhatnagar and Kallen, 1974; Bhatnagar and Meisami, 1998),
is noteworthy. Silver-stained extraepithelial neuronal
bodies also were described outside of the VNO in human
infants (Brookover, 1917). The significance of such ganglia is presently unclear. A possible explanation may lie
in putatively lost, aberrant LHRH neurons located outside typical migratory paths of vomeronasal or terminal
nerves described in mice (Wu et al., 1997). While clusters
of paravomeronasal neurons are common in fetal haplorhines that possess a VNO (Smith et al., 2003b), their
ubiquity in perinatal callitrichines is unique. At least
some callitrichines have a protracted period of postnatal
brain growth (Leigh, 2004). If a subset of the paravomeronasal cells is migratory, it is possible that their relatively
late occurrence is a reflection of other developmental
delays in the central nervous system.
Implications for Evolution of the VNS in
Platyrrhine Primates
Because of the complex role of olfactory communication in platyrrhines (Dobroruka, 1972; Epple, 1972;
1974; Milton, 1975; 1985; Boinski, 1987; Smith et al.
1997; Miller et al., 2003; Saltzman, 2003; Difiore et al.,
2006; Campos et al., 2007; Jones and Van Cantfort,
2007; Wolovich and Evans, 2007), this group of primates
presents a unique opportunity to understand the evolution of the VNS. Olfactory communication via scent
marking in platyrrhines is hypothesized to be important
for mediation of territoriality, social and/or reproductive
dominance, or intrasexual competition and mate attraction (Heymann, 2006), all of which have been hypothesized to be mediated by the VNS in other mammals (see
Halpern and Martı́nez-Marcos, 2003 for review). Since
genetic data suggest the common ancestor of platyrrhines had a functional VNS (Liman and Innan, 2003;
Zhang and Webb, 2003), there is reason to believe a sociosexual function played some part in the evolutionary
history of the platyrrhine VNS. The present study suggests this ancestral stock would eventually yield a great
morphological diversity, leaving much room for interpretation of its function in extant platyrrhines.
Our morphological survey of the platyrrhine VNO
indicates that some platyrrhines are similar to other
mammals (including at least some strepsirrhines) in certain respects. Notably, Ateles and both genera of marmosets have a VNO neuroepithelium with numerous rows
of VSNs. In addition, Callithrix and Ateles (but perhaps
not Cebuella) have terminally differentiated VSNs
within the VNNE, in keeping with observations on several rodents. Thus, Maier’s interpretation (1980) that
some platyrrhines exhibit ‘‘signs of reduction’’ (such as
the relatively thin VNNE and regions of poorly organized VNNE) does not denote a uniform characteristic of
platyrrhines. Maier’s statement is more evocative of
Saguinus spp, and perhaps Saimiri spp, those with the
thinnest VNNE in terms of nuclear rows (Smith et al.,
2004; and see Figs. 2 and 4) and also the smallest population of terminally differentiated VSNs (Dennis et al.,
2004; Smith et al., 2011). Although disagreements exist
about platyrrhine phylogenetic relationships (e.g.,
Steiper and Ruvolo, 2003; Ray et al., 2005; Hodgeson
et al., 2009; Rosenberger et al., 2009), those platyrrhines
with the thickest VNNE in terms of nuclear rows (marmosets and Ateles) or those that appear to have fewer
VSNs (Saguinus and Saimiri) are not regarded as especially close, as sister taxa (Fig. 14). Thus, no clear phylogenetic pattern is apparent. Our observations are
generally consistent with the assessment of Liman and
Innan (2003), who suggested selection pressure on a signal transduction channel related to VNO function is
reduced (but present) in all platyrrhines. Further, these
authors suggested ‘‘that in some species of the NW [New
World] monkeys the VNO may be vestigial or redundant’’ (p. 3331).
Further work on platyrrhines may clarify the persisting functional roles of the VNS, as well as the complex
manner in which sociosexual behaviors rely on the VNS
as opposed to other special senses. If some lineages of
platyrrhines are undergoing evolutionary regression of
the VNS, by what means do they detect social chemical
signals? In mammals, the MOS appears to have a synergistic overlap with the VNS (Powers and Winans, 1975;
Beauchamp et al., 1982). Thus it is unsurprising that
sociosexual functions that are strongly related to the
VNS of some mammals are taken up by the MOS in
taxa where the VNS is vestigial or absent (Wysocki and
Preti, 2004). If the MOS is mediating chemoreception in
some platyrrhines with reduced (if not vestigial) VNOs,
as in Saguinus spp, this may have implications for olfactory communication in other anthropoid primates like
Old World monkeys, apes, and humans. The results of
the present study isolate several lineages of living platyrrhines that may shed light on VNNE morphological
diversity (Fig. 14). In addition, our preliminary findings
suggest subtle osteological features, previously identified
on primates (Garrett et al., 2009), may be used to draw
inferences from fossil platyrrhines on the evolutionary
history of the VNS.
This study was made possible, in large part, by efforts
of numerous individuals who made well preserved tissues of rare specimens available to us over a ten year
period. A partial list of these individuals includes H.
Kafka and L. Gordon at the National Museum of Natural History, E. Less of the Cleveland Metroparks Zoo, E.
Curran of the New England Primate Center, L. Williams
at the Michael E. Keeling Center for Comparative Medicine and Research, J. Chatfield and T. deMaar of the
Gladys Porter Zoo, J. Trupkiewicz of the Philadelphia
Zoo, and S. Gibson of the University of South Alabama
Primate Research Center. In addition, authors thank all
other veterinary, research, or technical staff at these
institutions who took time to ensure that these specimens were preserved in formalin. Authors are grateful
to C. Vinyard and A. Taylor, who have shared cadaveric
tissue samples with them over the years. Some of the
specimens used in this study were sectioned by K. L.
Shimp and L. Maico-Tan.
Alport LJ. 2004. Comparative analysis of the role of olfaction and
the neocortex in primate intrasexual competition. Anat Rec A
Aujard F. 1997. Effect of vomeronasal organ removal on male sociosexual responses to female in a prosimian primate (Microcebus
murinus). Physiol Behav 62:1003–1008.
Barrett J, Abbott DH, George LM. 1993. Sensory cues and the suppression of reproduction in subordinate female marmoset monkeys, Callithrix jacchus. J Reprod Fertil 97:301–310.
Barton RA. 2006. Olfactory evolution and behavioral ecology in primates. Am J Primatol 68:545–558.
Beauchamp GK, Martin IG, Wysocki CJ, Wellington JL. 1982. Chemoinvestigatory and sexual behavior of male guinea pigs following vomeronasal organ removal. Physiol Behav 29:329–336.
Bhatnagar KP, Kallen FC. 1974. Morphology of the nasal cavities
and associated structures in Artibeus jamaicensis and Myotis lucifugus. Am J Anat 139:167–190.
Bhatnagar KP, Meisami E. 1998. Vomeronasal organ in bats and
primates: extremes of structural variability and its phylogenetic
implications. Microsc Res Technol 43:465–475.
Bhatnagar KP, Smith TD. 2006. The vomeronasal organ and its evolutionary loss in catarrhine primates. In: Preuss TM, Kaas JH,
editors. The evolution of primate nervous systems. Vol. 5. New
York: Elsevier. p 141–148.
Bhatnagar KP, Smith TD, Rodriguez-Duran A, Wible JR. 2006. The
Mormoopid vomeronasal organ (Chiroptera: Mormoopidae): observations on the VNO of Pteronotus macleayii and Pteronotus quadridens. Mammalia 70:288–292.
Boehm N, Gasser B. 1993. Sensory receptor-like cells in the human
foetal vomeronasal organ. NeuroReport 4:867–870.
Boinski S. 1987. Mating patterns in squirrel monkeys (Saimiri oerstedi). Behav Ecol Sociobiol 21:13–21.
Breipohl W, Bhatnagar KP, Mendoza A. 1979. Fine structure of the
receptor-free epithelium in the vomeronasal organ of the rat. Cell
Tissue Res 200:383–395.
Brennan PA. 2001. The vomeronasal system. Cell Mol Life Sci
Brookover C. 1917. The peripheral distribution of the nervus terminalis in an infant. J Comp Neurol 28:349–360.
Broom R. 1897. A contribution to the comparative anatomy of
the mammalian organ of Jacobson. Trans Roy Soc Edin 39:231–
Campos F, Manson JH, Perry S. 2007. Urine washing and sniffing
in wild white-faced capuchins (Cebus capucinus): testing functional hypotheses. Int J Primatol 28:55–72.
Cartmill M, Smith FH. 2009. The human lineage. Hoboken, NJ:
Cave AJE. 1973. The primate nasal fossa. Biol J Linn Soc London
Clancy AN, Coquelin A, Macrides F, Gorski RA, Noble EP. 1984.
Sexual behavior and aggression in male mice: involvement of the
vomeronasal system. J Neurosci 4:2222–2229.
Cooper JG, Bhatnagar KP. 1976. Comparative anatomy of the vomeronasal organ complex in bats. J Anat 122:571–601.
Dennis JC, Smith TD, Bhatnagar KP, Bonar CJ, Burrows AM, Morrison EE. 2004. Expression of neuron-specific markers by the
vomeronasal neuroepithelium in six species of primates. Anat Rec
Difiore A, Link A, Stephenson PA. 2006. Scent marking in two
Western Amazonian populations of woolly monkeys (Lagothrix
lagotricha). Am J Primatol 68:637–649.
Dobroruka L. 1972. Social communication in the brown capuchin:
Cebus apella. Int Zoo Yearb 12:43–45.
Doty RL. 2010. The great pheromone myth. Baltimore: Johns Hopkins University Press.
Epple G. 1972. Social communication by olfactory signals in marmosets. Int Zoo Yearbk 12:36–42.
Epple G. 1974. Olfactory communication in South American primates. Ann N Y Acad Sci 237:261–278.
Evans CS, Grigorieva EF. 1994. Morphology of the vomeronasal
organ in two South American monkeys (Saguinus labiatus and
Cebuella pygmaea, Callitrichidae): histology and lectin histochemistry. Adv Biosci 93:31–42.
Farbman AI, Margolis F. 1980. Olfactory marker protein during ontogeny: immunohistochemical localization. Dev Biol 74:205–215.
Frets GP. 1912. On the Jacobson’s organ in primates. Proc Sect Sci,
Akad Wetensch Amst 15:184–185.
Garrett EC. 2010. Getting in the groove: indirect observations of
the primate vomeronasal system using CT. Am J Phys Anthropol
Garrett EC, Smith TD, Burrows AM, Bonar CJ. 2009. Osteological
correlates of the vomeronasal system in primates. Am J Phys
Anthropol S48:132.
Gilad Y, Wiebe V, Przeworski M, Lancey D, Pääbo S. 2004. Loss of
olfactory receptor genes coincides with the acquisition of full trichromatic vision in primates. PLoS Biol 2:1–6.
Halpern M, Martı́nez-Marcos A. 2003. Structure and function of the
vomeronasal system: an update. Prog Neurobiol 70:245–318.
Hendrickx AG. 1971. Embryology of the baboon. Chicago: University of Chicago Press.
Hershkovitz P. 1977. Living New World monkeys (platyrrhini): with
an introduction to Primates. Chicago: University of Chicago
Heymann EW. 2006. The neglected sense—olfaction in primate
behavior, ecology, and evolution. Am J Primatol 68:519–524.
Hodgson JA, Sterner KN, Matthews LJ, Burrell AS, Jani RA,
Raaum RL, Stewart CB, Disotell TR. 2009. Successive radiations,
not stasis, in the South American primate fauna. Proc Natl Acad
Sci USA 106:5534–5539.
Hunter AJ, Fleming D, Dixon AF. 1984. The structure of the vomeronasal organ and nasopalatine ducts in Aotus trivirgatus and
some other primate species. J Anat 138:217–225.
Jackson LM, Harder JD. 1996. Vomeronasal organ removal blocks
pheromonal induction of estrus in gray short-tailed opossums
(Monodelphis domestica). Biol Reprod 54:506–512.
Jones CB, Van Cantfort TE. 2007. Multimodal communication by
male mantled howler monkeys (Alouatta palliata) in sexual contexts: a descriptive analysis. Folia Primatol 78:166–185.
Jordan J. 1972. The vomeronasal organ (of Jacobson) in primates.
Folia Morphol (Warsz) 31:418–432.
Kjær I, Fischer-Hansen B. 1996. Luteinizing hormone-releasing hormone and innervation pathways in human prenatal nasal submucosa: factors of importance in evaluating Kallmann’s syndrome.
APMIS 104:680–688.
Laska M, Rivas Bautista RM, Hernandez Salazar LT. 2005. Olfactory responsiveness to two odorous steroids in three species of
nonhuman primates. Chem Senses 30:505–511.
Laska M, Rivas Bautista RM, Hernandez Salazar LT. 2006. Olfactory sensitivity for aliphatic alcohols and aldehydes in spider
monkeys (Ateles geoffroyi). Am J Phys Anthropol 129:112–120.
Leigh SR. 2004. Brain growth, life history, and cognition in primate
and human evolution. Am J Primatol 62:139–164.
Lepri JJ, Wysocki CJ. 1987. Removal of the vomeronasal organ disrupts the activation of reproduction in female voles. Physiol
Behav 40:349–55.
Liman ER, Innan H. 2003. Relaxed selective pressure on an essential component of pheromone transduction in primate evolution.
Proc Natl Acad Sci USA 100:3328–3332.
Maier W. 1980. Nasal structures in old and New World primates.
In: Ciochon RL, Chiarelli AB, editors. Evolutionary biology of the
New World monkeys and continental drift. New York: Plenum
Press. p 219–241.
Matsui A, Go Y, Niimura Y. 2010. Degeneration of olfactory receptor
gene repertories in primates: no direct link to full trichromatic
vision. Mol Biol Evol 27:1192–1200.
Mendoza AS, Küderling I, Kuhn HJ, Kühnel W. 1994. The vomeronasal organ of the New World monkey Saguinus fuscicollis (Callitrichidae). A light and transmission electron microscopic study.
Ann Anat 176:217–222.
Meredith M, O’Connell RJ. 1979. Efferent control of stimulus access
to the hamster vomeronasal organ. J Physiol 286:301–316.
Miller KE, Laszlo K, Dietz JM. 2003. The role of scent marking in
the social communication of wild golden lion tamarins, Leontopithecus rosalia. Anim Behav 65:795–803.
Milton K. 1975. Urine-rubbing behavior in the mantled howler monkey (Alouatta palliata). Folia Primatol 23:105–112.
Milton K. 1985. Mating patterns of woolly spider monkeys, Brachyteles arachnoides: implications for female choice. Behav Ecol Sociobiol 17:53–59.
Monti-Graziadei AG. 1992. Cell migration from the olfactory neuroepithelium of neonatal and adult rodents. Dev Brain Res 70:65–74.
Nakajima T, Shiratori K, Ogawa K, Tanioka Y, Taniguchi K. 1998.
Lectin-binding patterns in the olfactory epithelium and vomeronasal organ of the common marmoset. J Vet Med Sci 60:1005–
Pestean A, Krizbai I, Bottcher H, Parducz A, Joo F, Wolff JR. 1995.
Identification of the Ulex europaeus agglutinin-I-binding protein
as a unique glycoform of the neural cell adhesion molecule in the
olfactory sensory axons of adult rats. Neurosci Lett 195:117–120.
Powers JB, SS Winans. 1975. Vomeronasal organ: critical role in
mediating sexual behavior of the male hamster. Science 187:961–
Ray DA, Xing J, Hedges DJ, Hall MA, Laborde ME, Anders BA,
White BR, Stoilova N, Fowlkes JD, Landry KE, Chemnick LG,
Ryder OA, Batzer MA. 2005. Alu insertion loci and platyrrhine
primate phylogeny. Mol Phylogen Evol 35:117–126.
Rosenberger AL, Tejedor MF, Cooke S, Halenar L, Pekkar S. 2009.
Platyrrhine ecophylogenetics, past and present. In: Garber P,
Estrada A, Bicca-Marques JC, Heymann, EW, Strier KB, editors.
South American primates: comparative perspectives in the study of
behavior, ecology and conservation. New York: Springer. p 69–113.
Roslinski DL, Bhatnagar KP, Burrows AM, Smith TD. 2000. Comparative morphology and histochemistry of glands associated with
the vomeronasal organ in humans, mouse lemurs, and voles.
Anat Rec 260:92–101.
Rossie JB. 2005. Anatomy of the nasal cavity and paranasal sinuses
in Aegyptopithecus and early Miocene African catarrhines. Am J
Phys Anthropol 126:250–267.
Rouquier S, Blancher A, Giorgi D. 2000. The olfactory receptor gene
repertoire in primates and mouse: evidence for reduction of the functional fraction in primates. Proc Natl Acad Sci USA 97:2870–2874.
Saltzman W. 2003. Reproductive competition among female common
marmosets (Callithrix jacchus): proximate and ultimate causes.
Proc Royal Soc B 276:197–229.
Salazar I, Sánchez Quinteiro P. 1998. Supporting tissue and vasculature of the mammalian vomeronasal organ: the rat as a model.
Microsc Res Technol 41:492–505.
Sánchez Villagra MR. 2001. Ontogenetic and phylogenetic transformations of the vomeronasal complex and nasal floor elements in
marsupial mammals. Zool J Linn Soc 131:459–479.
Satomi H, Takahashi K. 1986. The distribution and significance of
aberrant ganglion cells in the facial nerve trunk of the cat. Anat
Anz 162:41–46.
Shimp KL, Bhatnagar KP, Bonar CJ, Smith TD. 2003. Ontogeny of
the nasopalatine duct in primates. Anat Rec 274:862–869.
Smith TD, Bhatnagar KP. 2009. Vomeronasal system evolution. In:
Squire L, editor. New encyclopedia of neuroscience, Vol. 10.
Oxford: Academic Press. p 461–470.
Smith TD, Bhatnagar KP, Bonar CJ, Shimp KL, Mooney MP, Siegel
MI. 2003a. Ontogenetic characteristics of the vomeronasal organ
in Saguinus geoffroyi and Leontopithecus rosalia with comparisons to other primates. Am J Phys Anthropol 121:342–353.
Smith TD, Bhatnagar KP, Burrows AM, Shimp KL, Dennis JC,
Smith MA, Maico-Tan L, Morrison EE. 2005. The vomeronasal
organ of greater bushbabies (Otolemur spp.): Species, sex, and
age differences. J Neurocytol 34:135–147.
Smith TD, Bhatnagar KP, Shimp KL, Kinzinger JH, Bonar CJ, Burrows AM, Mooney MP, Siegel MI. 2002. Histological definition of
the vomeronasal organ in humans and chimpanzees with a comparison to other primates. Anat Rec 267:166–176.
Smith TD, Dennis JC, Bhatnagar KP, Bonar CJ, Burrows AM, Morrison EE. 2004. Ontogenetic observations on the vomeronasal
organ in two species of tamarins using neuron-specific b-tubulin
III. Anat Rec 278:409–418.
Smith TD, Dennis JC, Bhatnagar KP, Garrett EC, Bonar CJ, Morrison EE. 2011. Olfactory marker protein expression in the vomeronasal neuroepithelium of tamarins (Saguinus spp). Brain Res
Smith TD, Rossie JB. 2006. Primate olfaction: Anatomy and evolution. In: Brewer W, Castle D, Pantelis C, editors. Olfaction and
the brain: window to the mind. Cambridge University Press:
Cambridge. p 135–166.
Smith TD, Rossie JB, Bhatnagar KP. 2007. Evolution of the nose
and nasal skeleton in primates. Evol Anthropol 16:132–146.
Smith TD, Siegel MI, Bhatnagar KP. 2001a. Reappraisal of the vomeronasal system of catarrhine primates: ontogeny, morphology,
functionality, and persisting questions. Anat Rec 265:176–192.
Smith TD, Siegel MI, Bhatnagar KP. 2003b. Observations on the
vomeronasal organ of prenatal Tarsius bancanus borneanus with
implications for ancestral morphology. J Anat 203:473–481.
Smith TD, Siegel MI, Bonar CJ, Bhatnagar KP, Mooney MP, Burrows AM, Smith MA, Maico LM. 2001b. The existence of the vomeronasal organ in postnatal chimpanzees and evidence for its
homology to that of humans. J Anat 198:77–82.
Smith TE, Abbot DH, Tomlinson AJ, Mlotkiewicz JA. 1997. Differential display of investigative behavior permits discrimination of
scent signatures from familiar and unfamiliar socialldominant
female marmoset monkeys (Callithrix jacchus). J Chem Ecol
Starck D. 1975. Development of the chondrocranium in primates.
In: Luckett WP, Szalay FS, editors. Phyogeny of the primates.
New York: Plenum Press. p 127–155.
Steiper ME, Ruvulo M. 2003. New World monkey phylogeny based
on X-linked G6PD DNA sequences. Mol Phylogen Evol 27:121–
Stephan H, Frahm H, Baron G. 1981. New and revised data on volumes of brain structures in insectivores and primates. Folia Primatol 35:1–29.
Storan MJ, Key B. 2006. Septal organ of grüneberg is part of the olfactory system. J Comp Neurol 494:834–844.
Taniguchi K, Matsusaki Y, Ogawa K, Saito TR. 1992. Fine structure
of the vomeronasal organ in the common marmoset (Callithrix
jacchus). Folia Primatol 59:169–176.
Takagami S, Mori Y, Tanioka Y, Ichikawa M. 2004. Morphological
evidence for two types of mammalian vomeronasal system. Chem
Senses 29:301–310.
Tobet SA, Sower SA, Schwarting GA. 1997. Gonadotropin-releasing
hormone containing neurons and olfactory fibers during development: from lamprey to mammals. Brain Res Bull 44:479–486.
Wang G, Zhu Z, Shi P, Zhang Y. 2010. Comparative genomic analysis reveals more functional nasal chemoreceptors in nocturnal
mammals than diurnal mammals. Chin Sci Bull 55:3901–3910.
Weiler E, Benali A. 2005. Olfactory epithelia differentially express
neuronal markers. J Neurocytol 34:217–240.
Whinnett A, Mundy NI. 2003. Isolation of novel olfactory receptor
genes in marmosets (Callithrix): insights into pseudogene formation and evidence for functional degeneracy in non-human primates. Gene 304:87–96.
Wilson D, Hendrickx A. 1977. Quantitative aspects of proliferation
in the nasal epithelium of the rhesus monkey embryo. J Embryol
Exp Morphol 38:217–226.
Witt M, Georgiewa B, Knecht M, Hummel T. 2002. On the chemosensory nature of the vomeronasal epithelium in adult humans.
Histochem Cell Biol 117:493–509.
Wöhrmann-Repenning A. Bergmann M. 2001. The vomeronasal
complex in strepsirhine primates and Tarsius. Mamm Biol
Wolovich C K, Evans S. 2007. Sociosexual behavior and chemical
communication of Aotus nancymaae. Int J Primatol 28:1299–
Woodley SK, Cloe AL, Waters P, Baum MJ. 2004. Effects of vomeronasal organ removal on olfactory sex discrimination and odor
preferences of female ferrets. Chem Senses 29:659–669.
Wu TJ, Gibson MJ, Rogers MC, Sliverman AJ. 1997. New observations on the development of the gondaotropin-releasing system in
the mouse. J Neurobiol 33:983–998.
Wysocki CJ, Kruczek M, Wysocki LM, Lepri JJ. 1991. Activation of
reproduction in nulliparous and primiparous voles is blocked by
vomeronasal organ removal. Biol Reprod 45:611–616.
Wysocki CJ, Preti G. 2004. Facts, fallacies, fears and frustrations
with human pheromones. Anat Rec A 281:1201–1211.
Young JM, Massa HF, Hsu L, Trask BJ. 2010. Extreme variability
among mammalian V1R gene families. Genome Res 20:10–18.
Zhang J, Webb DM. 2003. Evolutionary deterioration of the vomeronasal pheromone transduction pathway in catarrhine primates.
Proc Natl Acad Sci USA 100:8337–8341.
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