peripheral sensory cells in the cephalic sensory organs of lymnaea stagnalis

Peripheral Sensory Cells in the Cephalic SensoryOrgans of Lymnaea stagnalis

Russell C. Wyeth1,2* and Roger P. Croll2

1Department of Biology, St. Francis Xavier University, Antigonish, Nova Scotia, B2G 2W5, Canada2Department of Physiology and Biophysics, Dalhousie University, Halifax, Nova Scotia, B3H 1X5, Canada

ABSTRACTThe peripheral nervous system in gastropods plays a

key role in the neural control of behaviors, but is poorly

studied in comparison with the central nervous system.

Peripheral sensory neurons, although known to be wide-

spread, have been studied in a patchwork fashion

across several species, with no comprehensive treat-

ment in any one species. We attempted to remedy this

limitation by cataloging peripheral sensory cells in the

cephalic sensory organs of Lymnaea stagnalis employing

backfills, vital stains, histochemistry, and immunohisto-

chemistry. By using at least two independent methods

to corroborate observations, we mapped four different

cell types. We have found two different populations of

bipolar sensory cells that appear to contain catecho-

lamines(s) and histamine, respectively. Each cell had a

peripheral soma, an epithelial process bearing cilia, and

a second process projecting to the central nervous sys-

tem. We also found evidence for two populations of ni-

tric oxide-producing sensory cells, one bipolar, probably

projecting centrally, and the second unipolar, with only

a single epithelial process and no axon. The various cell

types are presumably either mechanosensory or chemo-

sensory, but the complexity of their distributions does

not allow formation of hypotheses regarding modality.

In addition, our observations indicate that yet more

peripheral sensory cell types are present in the

cephalic sensory organs of L. stagnalis. These results

are an important step toward linking sensory cell mor-

phology to modality. Moreover, our observations

emphasize the size of the peripheral nervous system in

gastropods, and we suggest that greater emphasis be

placed on understanding its role in gastropod neuroe-

thology. J. Comp. Neurol. 519:1894–1913, 2011.

VC 2011 Wiley-Liss, Inc.

INDEXING TERMS: chemosensation; mechanosensation; peripheral nervous system; gastropod; Lymnaea

Gastropods have contributed greatly to our under-

standing of the neural control of behavior (Benjamin

et al., 2000; Chase, 2002; Elliott and Susswein, 2002;

Balaban, 2002; Getting, 2003; Crow and Tian, 2006;

Barco et al., 2006). Most of this work has explored cen-

tral mechanisms, including central pattern generators,

reflexive bases of behaviors, and the plasticity of those

reflexes. Accordingly, previous studies of identifiable sen-

sory neurons have focused on central sensory cells (e.g.,

Audesirk and Audesirk, 1980; Fredman and Jahan-Parwar,

1980; Nelson and Audesirk, 1986; Levenson et al., 2000;

Benjamin et al., 2000). However, peripheral sensory cells

(cells with peripheral somata and sensory endings) are

diverse, numerous, and presumably a major component

in the neural control of behavior (Lukowiak and Jacklet,

1972; Hoyle and Willows, 1973; Kupfermann et al., 1974;

Boudko et al., 1999; Croll et al., 2003; Leonard and

Edstrom, 2004). However, the majority of the physiologi-

cal experiments with peripheral sensory systems in gas-

tropods have examined responses mediated by unknown

cells (e.g., Janse, 1974; Bicker et al., 1982; Zaitseva

et al., 1987; Prescott et al., 1997; Murphy and Hadfield,

1997; Nakamura et al., 1999a; Kurokawa et al., 2008).

Neuroanatomical studies have described a number of

putative peripheral sensory cells (PSCs) in a range of gas-

tropod species. Histochemistry or immunohistochemistry

Additional Supporting Information may be found in the online version ofthis article.

Grant sponsor: Natural Sciences and Engineering Council of Canada (toR.C.W.); Grant sponsor: Natural Sciences and Engineering Council ofCanada; Grant number: Discovery grant 38863 (to R.P.C.); Grant sponsor:Malacological Society of London (to R.C.W.).

*CORRESPONDENCE TO: Russell Wyeth, Department of Biology, StFrancis Xavier University, P.O. Box 5000 Antigonish, NS B2G 2W5 Canada.E-mail: [email protected]

VC 2011 Wiley-Liss, Inc.

Received July 27, 2010; Revised October 11, 2010; Accepted January 17,2011

DOI 10.1002/cne.22607

Published online February 11, 2011 in Wiley Online Library(wileyonlinelibrary.com)

1894 The Journal of Comparative Neurology | Research in Systems Neuroscience 519:1894–1913 (2011)

RESEARCH ARTICLE

have been used to describe putative catecholaminergic

PSCs (Osborne and Cottrell, 1971; Salimova et al., 1987;

Croll et al., 1999; Voronezhskaya et al., 1999; Croll,

2001; Croll et al., 2003), histaminergic PSCs (Hegedus

et al., 2004), nitregic PSCs (Elphick et al., 1995; Serfozo

et al., 1998), FMRFamidergic PSCs (Nezlin et al.,

1994a,b; Suzuki et al., 1997; Wollesen et al., 2007; Faller

et al., 2008), and glutamatergic PSCs (Hatakeyama et al.,

2007). In addition, putative chemosensory cells have

been identified by using receptor signal transduction pro-

teins (Mazzatenta et al., 2004; Cummins et al., 2009).

Finally, electron microscopy and other methods have

been used to classify sensory cells based solely on struc-

tural features (e.g., Storch and Welsch, 1969; Zylstra,

1972; Kataoka, 1976; Emery and Audesirk, 1978; Davis

and Matera, 1982; reviewed in Emery, 1992; Steffensen

et al., 1993; Zaitseva, 1994; Nakamura et al., 1999b;

Gobbeler and Klussmann-Kolb, 2007).

However, with no comprehensive description in any

one species, the varied methods applied to a wide range

of species have greatly limited attempts to match PSC

types among the different studies. Consequently, our

goal here is to describe as many PSC types as possible in

one species, Lymnaea stagnalis, including morphology,

distributions, and putative neurotransmitter profiles.

These descriptions can both facilitate comparisons

among species and form the basis for future physiological

experimentation to assign modalities to the various

peripheral sensory cell types.

We chose to focus on the lips and tentacles (collec-

tively, the cephalic sensory organs [CSOs]; Fig. 1A) of the

pulmonate gastropod L. stagnalis for several reasons.

First, this species has been the subject of extensive prior

anatomical work (e.g., Zylstra, 1972; Zaitseva and

Bocharova, 1981; Croll and Chiasson, 1990; Elphick

et al., 1995; Croll et al., 1999; Hatakeyama et al., 2007),

thus providing the requisite background and tools for the

present study. Second, the CSOs likely contain both che-

mosensory and mechanosensory PSCs (de Vlieger, 1968;

Jager, 1971; Goldschmeding and Jager, 1973; Janse,

1976; Lever, 1977; Kemenes et al., 1986), thus providing

a variety of sensory cell types to be identified. Moreover,

examinations of feeding control in L. stagnalis have

explored both mechanosensation and chemosensation

mediated by the CSOs (Elphick et al., 1995; Staras et al.,

1999; Kemenes et al., 2001), adding physiological

relevance to our work. Finally, the thin and flat CSOs

(Fig. 1A) are suitable for whole-mount preparations, thus

allowing easy observation of PSC distributions as well as

morphology.

We first used general neuroanatomical methods to

label as many PSCs as possible in the CSOs, including

both backfills to label PSCs with axons in the CSO nerves,

and bath incubations in 1,10-dioctadecyl-3,3,30,30-tetra-

methylindocarbocyanine perchlorate (DiI), a lipophilic dye

Figure 1. Sampling locations for quantifying putative peripheral sensory cell distributions in cephalic sensory organs of Lymnaea stagnalis.

A: Anterior view of head, with the paired CSOs: lips (L) and tentacles (T). B: Schematic diagrams of isolated lips and tentacles (left CSOs,

ventral view) showing approximate locations of distal (black), intermediate (gray), and proximal sampling regions (outline), in medial (med),

central (cent; tip in tentacles), and lateral (lat) locations. In addition, a lateral strip region (hatched) was also sampled in tentacles.

Abbreviations

CSO cephalic sensory organFaGlu formaldehyde-glutaraldehydeHA histamineIR immunoreactiveNADPHd nicotinamide adenine dinucleotide phosphate diaphoraseNOS nitric oxide synthasePSC peripheral sensory cellTH tyrosine hydroxylase

Peripheral sensory cells in Lymnaea

The Journal of Comparative Neurology | Research in Systems Neuroscience 1895

that can accumulate preferentially in PSCs (Holland and

Yu, 2002). We then used neurotransmitter-specific

histochemistry and immunohistochemistry to distinguish

subsets of the PSCs, documenting morphologies, distri-

butions, and putative neurotransmitter content. We

report both greater morphological detail and distributions

for the catecholamine-containing PSCs, extend the

distribution for histamine-containing PSCs, and provide

evidence for at least two different populations of nitric

oxide-producing PSCs. The diversity of PSCs and their dis-

tributions emphasize the complexity of the peripheral

sensory nervous system in gastropods, and the need for

further study to understand the functional relevance of

that complexity in the control of behavior.

MATERIALS AND METHODS

AnimalsAll animal care and use was approved by the University

Committee on Laboratory Animals at Dalhousie Univer-

sity. A colony of Lymnaea stagnalis was maintained under

a 14–10-hour dark/light cycle at 26–28�C. Snails were

reared in a series of tanks with continuously aerated car-

bon-filtered tap water changed two or three times a week

and supplemented with yellow chalkboard chalk as a

hardener. Diet included both romaine lettuce and con-

sumer grade fish food (Nutrafin Max Sinking Pellets, Rolf

C. Hagen, Montreal, Quebec, Canada).

Dissection and fixation protocolThe lips and tentacles were dissected from snails (�10

mm shell length) in Lymnaea saline (LS; Wyeth et al.,

2009) with 0.125% 1-phenoxy-2-propanol (PP; Sigma, St.

Louis, MO, #484423) added as an anesthetic (Wyeth

et al., 2009). To improve the penetration of reagents into

peripheral tissues, we incubated CSOs for 15–90 minutes

in 0.25% collagenase (Sigma, #C9891) in PP-LS. After

rinsing (more than 2 minutes in PP-LS), the CSOs were

flattened between a glass coverslip and slide with model-

ing clay spacers at the corners (Wyeth et al., 2009), and

then held at 4�C for more than 15 minutes. CSOs were

fixed with 250 ll of 4�C fixative added slowly to one edge

of the coverslip so that fixative flowed past the tissues.

To complete fixation, CSOs were transferred to 1 ml of

4�C fixative. Dissected central nervous systems (CNSs)

were pinned in dishes lined in Sylgard (Dow-Corning, Mid-

land, MI), treated with 0.25% protease (Sigma, #P5147)

in LS for 5 minutes, and then rinsed in LS prior to fixation.

General neuroanatomical labelsBackfills

The peripheral projections of CSO nerves were back-

filled (n ¼ 15) by using methods from Croll (1986) and

Sakurai and Yamagishi (1998). The head was isolated in

PP-LS with CSOs and CNS still attached. The PP-LS was

replaced with LS, the target nerve was severed, and the

peripheral nerve root was sucked into a pulled glass micro-

pipette broken to match the nerve diameter. The micropip-

ette solution was then replaced with 5–25 ll of 4% Neuro-

biotin (Vector, Burlingame, CA, #SP1120) in distilled

water. After 12–36 hours at 4�C, the micropipette was

removed, and tissues were rinsed with LS. After a further

12–24 hours at 4�C, the CSOs were fixed overnight with

1% or 4% paraformaldehyde (PFA; Electron Microscopy Sci-

ences, Warrington, PA, #15710) in phosphate-buffered sa-

line (PBS; 0.1 M sodium phosphate with 0.9% sodium chlo-

ride, ph 7.4). Following fixation, tissues were rinsed in PBS

(four changes over 3 hours at room temperature on a

shaker), permeabilized for 4 hours with 4% Triton X-100 in

PBS (4% PBT), and incubated at 4�C for more than 24

hours in 10 lg/ml streptavidin Alexa Fluor 488 (Invitrogen,

Carlsbad, CA, #S11223) in 4% PBT. Tissues were then

rinsed in PBS, and mounted in a 3:1 mixture of glycerol

and 0.05 M Tris buffer at pH 8.0 with 2% propyl gallate

(Sigma, #P3130) added (Giloh and Sedat, 1982).

DiI vital stainFollowing Holland and Yu (2002), live L. stagnalis (n ¼

18) were incubated overnight in a 10-ml solution of fil-

tered water and 10 or 15 ll of 1 mg DiI (Invitrogen,

#D282) in 1 ml dimethyl formamide (Sigma, #D-4551).

CSOs were dissected and either mounted live or fixed

with 1% or 4% PFA in PBS overnight, rinsed and mounted

in PBS for viewing. (DiI labeling dissipated in glycerol.)

Neurotransmitter-specific labelsWe optimized all neurotransmitter-specific methods for

successful labeling throughout all regions of the CSOs.

Optimizations included different fixations, as well as

collagenase and detergent treatments to ensure reagent

penetration. Optimized methods for each label were used

for all observations and figures. However, we were unable

to similarly optimize double labels due to mutually exclu-

sive fixations or excessive tissue degradation during the

extra processing steps. For all neurotransmitter-specific

labels we also labeled CNSs to ensure our modified

methods reproduced previous results.

Basic immunohistochemistry protocolAll steps were conducted on a shaker at room temper-

ature or without shaking at 4�C, as noted. Following fixa-

tion with fresh fixatives, tissues were rinsed in PBS (four

changes over 3 hours), permeabilized for 4 hours with 4%

PBT, and blocked at 4�C overnight in 1% bovine serum

albumin (BSA; Sigma, #A4503) in 0.25% PBT (BSA-PBT).

Tissues were incubated at 4�C for 72 hours in primary

Wyeth and Croll

1896 The Journal of Comparative Neurology |Research in Systems Neuroscience

antibodies diluted in BSA-PBT, rinsed in 0.25% PBT, and

then blocked with 1% normal goat serum (Sigma,

#G9023) in 0.25% PBT (NGS-PBT). Tissues were next incu-

bated at 4�C for 48 hours in secondary antibodies diluted

in NGS-PBT, rinsed in PBS, and mounted in glycerol me-

dium. Secondary antibody specificities were verified in tis-

sues processed using identical procedures, except for the

elimination of the primary antibodies (Table 1).

Anti-tyrosine hydroxylaseimmunohistochemistry

Tyrosine hydroxylase (TH) is involved in the production

of dopamine and other catecholamines in molluscs (Pani

and Croll, 1998), and anti-TH immunohistochemistry has

been used previously to label putative catecholaminergic

cells in L. stagnalis (Croll and Chiasson, 1990; Croll et al.,

1999). Tissues were fixed, including the initial 1 hour

under a coverslip and subsequently in a vial, in either 4%

PFA in 0.1 M PBS for 3 hours total or 1% PFA overnight.

Anti-TH primary antibodies (Table 2) were labeled with

goat anti-mouse AlexaFluor 555 secondary antibodies

(Invitrogen, #A21422; diluted 1:100). TH immunoreactiv-

ity in the CNS was similar to a previous description using

the same antibody (Croll et al., 1999).

FaGlu histochemistryWe used the formaldehyde-glutaraldehyde (FaGlu)

method as a second label for catecholamine-containing

cells (Furness et al., 1978). Tissues were fixed overnight

in a 4% PFA and 0.5% glutaraldehyde (Electron

Microscopy Sciences, 16320) in 0.1 M PBS. Blue-green

fluorescence under UV illumination, indicative of cate-

cholamines, was observed after the tissues were rinsed

twice in distilled water and air-dried overnight (Hauser

and Koopowitz, 1987), and mounted in methyl salicylate.

Control tissues were fixed without glutaraldehyde, and

showed no corresponding fluorescence.

Anti-histamine immunohistochemistryTissues were fixed in 2% 1-ethyl-3(3-dimethyl-amino-

propyl)-carbodiimide (EDAC; Sigma, #E7750) with 0.4%

n-hydroxysuccinimide (Sigma, #H7377) for 3 hours (1

hour under coverslip, 2 hours in a vial), followed by 1%

PFA overnight (Panula et al., 1988). Nonspecific label

was minimized when tissues and primary antibodies

were preincubated separately in BSA-PBT with 200 lg/ml keyhole limpet hemocyanin (Sigma, #H5654). Alter-

natively, similar but less intense labeling was also

found in some tissues fixed with 1% PFA overnight.

Rabbit anti-histamine (HA) antibodies (Table 2) were la-

beled with anti-rabbit AlexaFluor 488 secondary anti-

bodies (Invitrogen, #A11008; diluted 1:100). HA immu-

noreactivity in the CNS was similar to a previous

description using the same antibody (Hegedus et al.,

2004).

TABLE 2.

Primary Antibodies Used to Label Putative Peripheral Sensory Cells in Lymnaea stagnalis

Antibody Immunogen Details Dilution

Tyrosine hydroxylase (TH) TH from rat PC12 cells; epitope in widelyconserved mid-portion

Immunostar 22941 mouse, monoclonal 1:250

Histamine (HA) HA conjugated to succinylated keyholelimpet hemocyanin by carbodiimide

Immunostar 22939 rabbit, polyclonal 1:250

Nitric oxide synthase (NOS) Peptide: DQKRYHEDIFG (‘‘uNOS’’—sequenceis conserved among NOS isoforms)

Affinity Bioreagents PA1-039rabbit, polyclonal

1:100

TABLE 1.

Sample Sizes for Protocols Used to Label Putative Peripheral Sensory Cells (PSCs) in Lymnaea stagnalis Sensory Organs1

Immunohistochemistry Corroboration

Protocol Positive Preincubation control Secondary control Protocol Positive Control

Anti-TH 27 NA 3 FaGlu 13 12

Anti-HA 14 3 3 NAAnti-NOS3 Uni: 7 NA 3 NADPHd 20 NADP: 5

Uni & bi: 8 NADH: 7NAD: 3

1Number of snails with positive labeling using immunohistochemistry protocols for PSCs, along with preincubation controls controls, and secondary

controls without primary antibodies. Sample sizes are also given for histochemical protocols and their corresponding controls used to corroborate

the immunohistochemistry. NA, not available. For abbreviations, see list.2Although only one dedicated FaGlu control was performed here, all paraformaldehyde-fixed tissues also serve as controls3Anti-NOS samples sizes for animals with only unipolar (Uni) or both unipolar and bipolar (Uni & bi) PSCs.



Anti-nitric oxide synthaseimmunohistochemistry

Nitric oxide synthase (NOS) is an enzyme that pro-

duces nitric oxide, and other anti-NOS antibodies have

been used previously to identify putative nitrergic cells

(Dawson et al., 1991; Moroz et al., 1999). Tissues were

fixed in 1% PFA overnight. Rabbit anti-NOS primary anti-

bodies (Table 2) were labeled with goat anti-rabbit Alexa-

Fluor 488 secondary antibodies (Invitrogen, #A11008;

diluted 1:100).

NADPH diaphorase histochemistryWe used NADPH diaphorase (NADPHd) histochemistry

as a second label for NOS-containing cells (Scherersin-

gler et al., 1983; Dawson et al., 1991; Elofsson et al.,

1993; Moroz et al., 1999), This method has been sug-

gested as the best option for identifying putative nitrergic

cells in gastropods (Moroz, 2000, 2006). Following the

protocol of Ott and Elphick (2003), we fixed CSOs with an

ice-cold 9:1 methanol/formalin mixture (1 hour under a

coverslip, followed by 1–2 hours in a vial), rinsed in Tris

buffer (TB), pH 7.2, and permeabilized in 0.2% Triton-X in

TB, pH 7.2. The tissues were incubated for 3 hours in the

dark in a reagent mixture (1 mM b-NADPH) and 0.5 mM

nitro blue tetrazolium in 0.2% Triton-X in TB, pH 8) in

ice-cold conditions, before incubation overnight at room

temperature in the dark. Following rinsing in TB, pH

7.2, and dehydration through an ethanol series, nonspe-

cific overstain was removed by treatment with 100%

methanol (three 15 min changes), before clearing and

mounting in cedar oil.

Specificity controls were performed by substituting b-NADP, NADH, and NAD for NADPH in the reagent mixture.

Both NAD and NADP and four NADH controls showed no

labeling. However, three trials with NADH showed both

peripheral and central labeling, similar to tissues labeled

simultaneously with b-NADPH, albeit less intense and lessextensive. We hypothesize that our long incubations, used

to achieve labeling in the peripheral tissues, allowed a low

reaction rate with NADH to still produce positive label.

Previous reports from L. stagnalis either did not use

NADH as a control (Serfozo et al., 1998) or used much

shorter reagent incubations (Moroz et al., 1993; Moroz

et al., 1994a). Furthermore, we observed NADPHd reactiv-

ity in the CNS consistent with previous reports describing

putative central nitrergic cells in L. stagnalis (Moroz et al.,

1993; Moroz et al., 1994a,b; Serfozo et al., 1998). Thus,

despite the ambiguous control results, we suggest that

our NADPHd labeling was specific for NOS.

Antibody specificitiesAnti-TH specificity was supported by our observations

that TH immunoreactivity in both the CNS and periphery

was similar to catecholamine histochemical labeling with

the FaGlu method, as has also been shown previously

(Croll and Chiasson, 1990; Croll et al., 1999). Anti-HA

specificity was supported by the absence of labeled PSCs

and nerves in our observations of tissues processed with

primary antibodies preadsorbed with 40 lg/ml histamine

conjugated to BSA by EDAC (Panula et al., 1984, 1988).

Preincubation did not affect labeling of other structures

(most notably the superficial surface of some epithelial

cells), which we therefore excluded from analysis. The

evidence for anti-NOS specificity using NADPHd as a

second label for NOS is less clear. We observed similar

NOS immunoreactivity and NADPHd reactivity in the

CNS, particularly a ring of fibers through the neuropils of

the cerebral and visceral loop ganglia, as well as proc-

esses in the CSO nerves. However, in the periphery, we

found structures that matched using the two labeling

methods (processes in the nerves branching throughout

the CSO and unipolar PSCs), as well as substantical dif-

ferences (bipolar PSCs found only in observations of NOS

immunoreactivity), and our conclusions are tempered

accordingly.

Double labelsDouble-labeling procedures pairing backfills, DiI, fix-

able analogs of DiI (Invitrogen #L7781 and #C7001), and

all anti-sera with different primary hosts were attempted

by using a range of compromises between the various

protocols.

ObservationsLaser scanning confocal image stacks of fluorescent

immunohistochemical labeling were acquired by using

Zeiss LSM 510 or 510 Meta microscopes and Zeiss LSM

software (Carl Zeiss, Thornwood, NY). Epifluorescence

micrographs of FaGlu labeling were acquired and other

fluorescent labels were observed by using a Leica

DM4000B microscope equipped with D, L5, and N2.1 fil-

ter sets and a Leica DC500 camera and Leica Application

Suite (v2) software (Leica, Deerfield, IL). Transmitted light

micrographs of NADPH diaphorase labeling were

acquired by using an Axioplan 2 microscope equipped

with a Axiocam HRc digital camera and an AxioVision

v4.6 software (Carl Zeiss). Images were processed by

using CombineZP (Alan Hadley), ImageJ (National Insti-

tutes of Health), ZEN 2008, and 2009 LE (Carl Zeiss),

Matlab R14 (Mathworks, Natick, MA), or Photoshop v7

(Adobe Systems, San Jose, CA). Processing included

image-wide intensity adjustments, median filters to

improve clarity for anti-NOS images, and cropping.

Images in figures are either maximum projections of

confocal image stacks, minimum projections of multiple

brightfield focal planes (grayscale brightfield stacks were

Wyeth and Croll


inverted, maximum projected, and the projection inverted

again), focused stacks of brightfield focal planes (Stack

Focuser plugin in ImageJ), combined stacks of epifluores-

cence focal planes (default settings in CombineZP), or

single brightfield focal planes. Multiple adjacent images

were joined together with ImageJ and the TurboReg and

MosaicJ plugins (Thevenaz and Unser, 2007). Schematics

were created in Illustrator v10 (Adobe Systems), and fig-

ures were assembled and labeled with InDesign v2

(Adobe Systems)

QuantificationPSCs were measured in at least six CSOs by using an

Olympus BX50 microscope equipped with an Optronics

Microfire digital camera, and Optiscan motorized stage

(Prior Scientific, Rockland, MA), and Neurolucida 7.5 soft-

ware (MBF Bioscience, Williston, VT). Epithelial process

lengths were measured by tracing in three dimensions,

and somata diameters were approximated by the maxi-

mum feret of outlines traced around the somata. We could

not compensate for different changes in tissue size result-

ing from the different labeling methods (in particular,

NADPH diaphorase caused tissues to shrink more than the

other methods). Both lengths and diameters were com-

pared separately by using ANOVAs, followed by Tukey’s

post hoc comparisons (a ¼ 0.05) in SPSS v15 (SPSS, Chi-

cago, IL). Anti-NOS-reactive cells were measured in confo-

cal stacks because other structures obscured the sensory

cells during epifluorescence observation.

PSC densities were quantified with Neurolucida soft-

ware by counting cells inside 150 � 150-lm sampling

regions. In the lips, we sampled six regions for both dorsal

and ventral surfaces (Fig. 1B). The regions were placed

systematically along the lip margin: medial at 17% of the

margin length, central at 50%, and lateral at 83%. For

each of these locations, distal regions were placed

directly tangential to the lip margin, and proximal regions

300 lm along a line perpendicular to the margin. In the

tentacles, regions were placed at the tip and 150 lmproximal to the tip, as well as at 50% of the medial margin

and 50% of the lateral margin, for both dorsal and ventral

surfaces. For histamine-immunoreactive PSCs, an addi-

tional region was placed over the high-density strip of

PSCs inset from the lateral margin. For NADPHd-reactive

PSCs, intermediate regions at 150 lm from the margin

were used in the lip, and also in the medial and lateral

tentacle. PSCs were only counted if both soma and den-

drite were inside the region.

RESULTS

We focused our observations on the CSOs of L. stagna-

lis, identifying unipolar PSCs with a putative sensory

dendrite that penetrated the epithelium, and also bipolar

PSCs, with a second process extending elsewhere in the

CSO tissue. By using these criteria, we were able to verify

distributions (summarized in Fig. 2) and morphologies for

Figure 2. Schematic diagram of the relative distributions of puta-

tive peripheral sensory cells in the cephalic sensory organs of L.

stagnalis. Shading indicates PSC densities: high (dark gray), inter-

mediate (light gray), low (white). A: Explanatory map showing how

both dorsal (d) and ventral (v) surfaces of the lips (l) and ten-

tacles (t) with medial (m) and lateral (lat) regions are displayed,

and their relationship to the eyes (filled circles) and mouth (filled

semicircle). B: TH-IR bipolar PSCs. C: HA-IR bipolar PSCs. D:

NOS-IR bipolar PSCs, for whom the distribution is tentative (?)

given the capricious nature of the immunoreactivity. E: NADPHd-R

unipolar PSCs (also observed with anti-NOS labeling). For other

abbreviations, see list.



four types of PSCs by using two or more labeling methods

in each case (Table 3).

General neuroanatomical labelsBackfills

We first attempted to label all bipolar PSCs by using

backfills of the nerves leading to the CSOs. This method

labeled an extensive peripheral nervous system

(Fig. 3A,B). Numerous PSCs were present, with at least

three qualitatively recognizable types, distinguishable

based on the length and shape of their epithelial process,

the degree to which their somata formed clusters, and

their distribution in tentacles (Fig. 3C–H). We also

observed systematic differences in the frequency of label-

ing with one cell type labeled in all backfills whereas

others labeled only when filling appeared to be more com-

plete. The first type of PSCs, observed in all backfills, was

solitary (i.e., with somata that did not form distinct clus-

ters) and had relatively long epithelial processes (�40

lm; Figs. 3C,E, 4A) that often extended parallel to the

epithelial surface before bending and penetrating the epi-

thelium (Fig 3D; Supplementary Movie 1). These cells

were found primarily along a ventral lateral strip of the

tentacles, just medial to the lateral margin (n ¼ 9) and

also in the lips (n¼ 4).

A second population of single PSCs, observed in most

preparations, had relatively shorter epithelial processes

(�20 lm; Figs. 3C, 4B) and was found along the margins

of both the tentacles (n ¼ 6) and lips (n ¼ 5). The epithe-

lial processes in these PSCs did not run parallel to the

epithelial surface, and thus lacked the distinct bend in

the process. Finally, in preparations with the most com-

plete labeling, PSCs with clustered somata and relatively

short epithelial processes also without a distinct bend

(Figs. 3F–H, 4C; Supplementary Movie 2) were found in a

ventral patch on the tentacles (n ¼ 7; Fig. 3B), as well as

the proximal medial margin (Fig. 3C) and other areas of

the tentacle. We attribute the differences in labeling reli-

ability for the three PSC types to either differing axonal

transport rates or axon closure rates after the nerve was

severed prior to backfilling, both possibly correlated with

axon diameter. Cilia were present on at least some exam-

ples of all three sensory cell types. Other backfilled PSCs

were present beyond those noted here, but the density of

neural structures with the most complete labeling made

interpretation difficult (Fig. 3I). We have therefore limited

our description to PSCs with morphology and distribu-

tions consistent with those also labeled by more specific

methods.

TABLE 3.

Hypothesized Neurotransmitters and Matching Between

Different Labeling Protocols for Putative Peripheral

Sensory Cells (PSCs) in the Cephalic Sensory

Organs of Lymnaea stagnalis1

PSC type IHC Histochemistry DiI Backfill

Catecholaminergic:bipolar

a-TH FaGlu Dim Single, short

Histaminergic:bipolar

a-HA NA Bright Single,long, bent

Nitrergic: unipolar a-NOS NADPHd — NANitrergic: bipolar a-NOS — — Clustered

PSCs were labeled using immunohistochemistry (IHC), histochemistry

to corroborate the IHC, DiI vital stain, which labeled PSCs consistent

with a-TH PSCs (dim labeling) and a-HA PSCs (bright labeling), and

backfills, which labeled PSCs with similar morphologies and distribu-

tions to all three types of bipolar PSCs (with either single or clustered

somata and shorter or longer and bent dendrites). NA, no applicable

labeling method; �, no label observed. For other abbreviations,

see list.

Figure 3. Backfills label at least three types of putative PSCs in the CSOs of L. stagnalis. A: Dorsal view of a backfilled tentacle demon-

strates the extensive innervation of the CSOs, including dense areas of PSCs along the medial margin (m) and around the eye (e). B: Ven-

tral view of a backfilled tentacle (without the eye) shows a ventral patch of high-density PSCs (p) and an additional lateral strip of PSCs

(s). C: Higher magnification of a tentacle medial margin reveals three morphologically distinct subepithelial bipolar PSC types: type 1

(open arrowheads), solitary (i.e., not in clusters), with a longer and often bent (b) epithelial process, and found primarily along the lateral

ventral strip; type 2 (arrowheads), solitary, with relatively short epithelial processes; and type 3 (double arrowheads), clustered, with rela-

tively straight epithelial processes. Supplementary Movie 1 scans through the confocal stack used for this projection, and provides an

alternate view of the three PSC types. D: Orthogonal projection of region outlined by dashed lines in C illustrates the difference between

type 1 PSCs (open arrowhead) with an epithelial process that ran parallel to the sensory epithelium before bending (b) and extending

through the epithelium and type 2 PSCs (arrowhead) with processes that extend directly through the epithelium. E: In addition, the first

type of PSC (four shown here from a ventral view of a tentacle) were sometimes the only PSCs labeled. F: The clustered PSCs formed the

ventral patch, with very high densities and uniformly distributed epithelial processes (see Supplementary Movie 2 for an animated view of

this confocal image series). G: A single optical slice 12 lm below the epithelial surface (same confocal image series as projected in E)

shows that the epithelial processes had a relatively uniform distribution. H: A single optical slice 27 lm below the epithelial surface (same

confocal image series as projected in E) shows tightly clustered somata (arrowheads) separated by regions with no backfill label (open

arrowhead). I: A 73 � 73 � 42-lm-deep region of the lateral margin of tentacle has at least 50 PSCs that are not easily classified among

the three types. Scale bar ¼ 500 lm in A,B; 50 lm in C,E,F,G,H; 20 lm in D,I.

Wyeth and Croll


DiITwo types of bipolar PSCs were labeled in CSOs of live

snails immersed in DiI solutions. The first type labeled

more strongly (Fig. 5A,B) and had, on average, 34-lm-

long epithelial processes (Table 4). Sometimes a single

cilium (or tight cluster of cilia) extended from the tip of

the process beyond the epithelium surface (Fig. 5E).

These PSCs had a limited distribution, found primarily on

a strip along the ventral surface of the tentacle (Fig. 6)

and the lateral margins of the lips. The second showed

much weaker (and rapidly fading) fluorescence (Fig.

5A,B), making exact measurements unreliable. In live

preparations, a tuft of motile cilia was sometimes visible

at the end of the process extending beyond the

Figure 3



epithelium (Fig. 5D; Supplementary Movie 3). These PSCs

were more widespread, but were erratically labeled, rang-

ing from only a few cells to more than 100 per tentacle,

and they were particularly dense along the medial margin.

Both cell types had processes that joined the nerves that

branch throughout the CSOs (Fig. 5B) and at least one

and possibly both extended into the CNS, as DiI fluores-

cence was also observable in the CSO nerves and

cerebral ganglia (Fig. 5C). Thus, these two bipolar PSCs

labeled with DiI appeared to match the two solitary cell

types in backfills (i.e., those that did not have clustered

somata). No other cells consistent with either a unipolar

or bipolar sensory morphology were labeled by DiI.

Neurotransmitter-specific labelsWe next used a combination of immunohistochemistry

and (where possible) a second histochemical label to

further explore PSC populations in the CSOs. These more

specific methods reduced the complexity of labeling, and

thus allowed more precise descriptions of distributions

and morphologies.

Anti-tyrosine hydroxylase and FaGluNumerous TH-immunoreactive (IR) PSCs were

observed in the tentacles and lips of L. stagnalis (Fig. 7).

The cells had 23 6 7-lm-long epithelial processes (mean

6 SD; Table 4), and some appeared to be ciliated (Fig.

7A). The TH-IR PSCs were especially dense on the ventral

medial surface of the lip, along the entire margin of the

lip, and on the medial margin and tips of the tentacles

(Figs. 2B, 7B,C, 8A). Elsewhere, the densities were lower,

Figure 4. Schematic diagram demonstrating the difference

between the three types of bipolar PSCs labelled in backfills.

A: The first type had solitary somata with longer epithelial proc-

esses (�40 lm) that extended parallel to the epithelial surface

with a distinct bend before penetrating the epithelium. B: The

second type had solitary somata and shorter epithelial processes

(�20 lm) without the distinct bend. C: The third type had clus-

tered somata and shorter epithelial processes (�20 lm) without

a distinct bend. Scale bar ¼ 10 lm in A.

Figure 5. DiI labels two types of putative PSCs in the CSOs of L. stagnalis. A: Both dim (d) and bright (b) bipolar PSCs had processes

(arrowheads) that penetrate the tentacle epithelium and another (arrows) that joined the branching innervation of the CSOs. Live tissue.

B: The branching innervation of a tentacle includes processes (arrows) from numerous dim (d) or bright (b) (or both) DiI PSCs. Live tissue.

C: These processes were apparent centrally, in the tentacle nerve (tn) and penetrated the neuropil (arrow) of the cerebral ganglion. Fixed

tissue. D: The dimmer PSCs bore a tuft of cilia (*) that were sometimes motile in live tissue (see Supplementary Movie 3). E: The brighter

PSCs also bore a single large cilium (or tight cluster of cilia) that were never motile. Fixed tissue. Scale bar ¼ 20 lm in A,D,E; 50 lm in

B; 100 lm in C.

Wyeth and Croll


particularly along the lateral edge of the tentacles (Fig.

8A). The TH-IR PSCs were consistently bipolar, and the

branched nerves of the CSOs were prominently TH-IR

(Fig. 7C); however, the high density of labeled fibers

made it difficult to determine what proportion of PSC

processes continued in the nerves and thus contributed

to the substantial TH immunoreactivity in the CSO nerves

as they entered the ganglia of the CNS (data not shown;

Croll and Chiasson, 1990; Croll et al., 1999). We also

observed TH-IR peripheral neurons without dendrites, as

well as a rectilinear pattern of TH-IR fibers in areas with

and without TH-IR PSCs in both CSOs (Fig. 7D). Overall,

the population of TH-IR PSCs had characteristics most

similar to the PSCs dimly labeled by DiI and the solitary

PSC type with shorter dendrites in backfills (Table 3).

We corroborated our observations of TH-IR PSCs with

observations of a similar population of blue-green fluores-

cent PSCs in FaGlu-fixed CSOs (Fig. 9). FaGlu-labeled

PSCs had an epithelial process (Fig. 9A), were bipolar,

and were found particularly along the CSO margins (Fig.

9B,C). In addition, the relative size of the epithelial pro-

cess and soma in these cells was qualitatively similar to

that of the TH-IR PSCs. Thus, we suggest that the same

PSCs were labeled by using both FaGlu and anti-TH.

Anti-histamineWe observed many HA-IR PSCs in L. stagnalis CSOs

(Fig. 10). The cells had, on average, 426 10-lm-long epi-

thelial processes without labeled cilia (Fig. 10A, Table 4).

HA-IR PSCs were found along a ventral strip in the ten-

tacles and over much of the ventral surface of the lips,

but were scarce in other CSO regions (Figs. 2C, 8B, 10B–

D). The cells were consistently bipolar, and their proc-

esses apparently joined the branched CSOs nerves (Fig.

10C,D) and presumably contributed to the HA-IR fibers

present in all three CSO nerves as they entered the CNS

(data not shown). The distribution and morphology of

these PSCs were thus similar to both the PSCs brightly la-

beled by DiI and the single PSCs with longer dendrites in

backfills (Table 3).

In addition to the PSCs, a number of HA-IR fibers with-

out associated somata were also present, extending from

the major nerves branching in the CSOs (Fig. 10C,D). The

fibers were distributed similarly to the HA-IR PSCs in the

tentacles, but in some preparations were found through-

out the lips, not just laterally, as were the PSCs.

Anti-nitric oxide synthase and NADPHdiaphorase

We used both anti-NOS immunohistochemistry and

NADPH diaphorase histochemistry as independent meth-

ods to identify putative nitrergic PSCs. NOS immunoreac-

tivity was inconsistent. In approximately half the prepara-

tions, numerous bipolar PSCs were present (Table 1, Fig.

11A,B). Although not quantifiable in comparable fashion

to the other PSC types, these cells had clustered somata

and 44 6 18-lm-long epithelial processes (Table 4) with-

out labeled cilia. These cells were observed along the

medial margin, tip, and a ventral strip of the tentacles,

although distributions varied among preparations (Fig.

2D). Processes from the bipolar NOS-IR PSCs apparently

joined the branched nerves of the CSOs, and NOS immu-

noreactivity was observed in all three CSO nerves as they

entered the CNS (data not shown). Thus, there were

some morphological similarities between these cells and

clustered bipolar PSCs observed in backfills, but the

backfilled PSCs were more widespread than the cumula-

tive distribution of bipolar NOS-IR PSCs observed across

all preparations.

TABLE 4.

Dendrite Lengths and Soma Diameters of Putative

Peripheral Sensory Cells in the Cephalic Sensory Organs

of Lymnaea stagnalis Labeled by Using Four Different

Methods1

Label Cell type

Dendrites

(lm)

Somata

(lm) No.

DiI: bright cells Bipolar 34 6 12* 11 6 2* 50a-TH Bipolar 23 6 7** 11 6 2* 70a-HA Bipolar 42 6 10*** 11 6 2* 62NADPHd Monopolar 11 6 3**** 9 6 3** 60a-NOS2 Bipolar 44 6 18 9 6 2 10

The methods used were DiI vital stain, anti-tyrosine hydroxylase

immunohistochemistry (a-TH), anti-histamine immunohistochemistry

(a-HA), NADPH diaphorase histochemistry (NADPHd), and anti-nitric ox-

ide immunohistochemistry (a-NOS). The data are means 6 SD. For den-

drites, significantly different lengths between labelled PSCs, based on an

ANOVA followed by Tukey’s post-hoc comparisons a ¼ 0.05), are indi-

cated by the different numbers of asterisks. For somata, significantly dif-

ferent sizes are similarly indicated by the different numbers of asterisks.2Measured from confocal stacks, not using Neurolucida software as were

the other PSC types, and thus not part of the statistical comparison.

Figure 6. Densities of bright DiI putative peripheral sensory cells

(PSCs) in the cephalic sensory organs of L. stagnalis. Inset shows

where sensory cells were counted in the medial, tip, lateral, and

lateral strip (hatched) regions of the tentacles (orientations: Med,

medial; Prox, proximal). Numbers in parentheses indicate sample

sizes.



A second population of unipolar NOS-IR cells with only

a single epithelial process was often observed (Fig. 11A),

and similar cells were consistently labeled by using

NADPH diaphorase (Fig. 11C,D). The NADPHd-reactive

(NADPHd-R) cells (Fig. 11D) had 11 6 3-lm-long epithe-

lial processes (Table 4), and none appeared to be ciliated.

The epithelial process diameters (1–3 lm) appeared to

be larger than those of the other PSC types (1 lm). The

NADPHd-R PSCs were found on all surfaces of the CSOs,

with highest densities on the dorsal lip surface (Figs. 2E,

8C). These PSCs were consistently unipolar, with no

connection to NADPHd-R label in the branched nerves of

the CSOs (Fig. 11D). No bipolar NADPHd-R counterparts

to the NOS-IR bipolar PSCs were observed. Overall, these

observations suggest two populations of PSCs, one

bipolar and the other unipolar.

Double labeling and size comparisonsTo test whether the various PSCs were separate popu-

lations, we attempted double labels, pairing different anti-

bodies as well as antibodies with backfills. Pairing anti-TH

and anti-HA revealed no overlap in immunoreactivity, and

a few PSCs could be backfilled and labeled with either

anti-TH or anti-HA. However, the tissues were noticeably

degraded after these longer protocols, and we were

Figure 7. Anti-tyrosine hydroxylase immunoreactive bipolar putative PSCs in the CSOs of L. stagnalis. A: TH-IR PSCs had one process that

penetrated the epithelium (arrowhead) and another (arrow) that joined the branching innervation of the CSOs. Inset: the same confocal

stack, adjusted for high contrast, overlaid with a brightfield image acquired at the same time suggests that the TH-IR PSCs bear a tuft of

cilia (open arrowhead in both views) that extend beyond the epithelial surface (dotted line). B: The TH-IR PSCs were concentrated particu-

larly over the ventral medial surface (dashed line) and margin of the lip (arrowheads), with only the deeper branching innervation (arrows)

apparent in lateral regions. C: The medial margin of the tentacles also showed a high density of TH-IR PSCs, with at least some their proc-

esses joining the branching innervation (arrows) of the CSO. D: The lateral tentacle had few TH-IR PSCs, making the approximately rectilin-

ear pattern of TH-IR processes the most prominent feature. Scale bar ¼ 5 lm in A and inset; 400 lm in B; 50 lm in C,D.

Wyeth and Croll


unable to replicate the results for any label in isolation.

Double labels involving either the NOS antibody DiI (or its

fixable analogs) did not provide satisfactory labeling of

PSCs. In the absence of consistent double labeling, we

used size comparisons among the different PSC popula-

tions. Epithelial process lengths for each testable cell

type were significantly different from all other cell types

(ANOVA, F3,238 ¼ 157, P < 0.001). Process lengths, in

order from shortest to longest were as follows (Tukey’s

post hoc comparisons): NADPHd-R PSCs < TH-IR PSCs <

Figure 8. Putative PSC densities in the cephalic sensory organs of L. stagnalis. A: Anti-tyrosine hydroxylase immunoreactive PSCs. B: Anti-

histamine immunoreactive PSCs. C: NADPH diaphorase reactive unipolar PSCs. Insets show where sensory cells were counted in distal,

(black bars) intermediate (gray bars), or proximal (white bars) locations in the medial (med), central (cent), and lateral (lat) regions of the

lips, or medial, tip, and lateral regions of the tentacles. For HA-IR PSCs, an additional lateral strip region was also counted (hatched bars).

Orientations: Med, medial; Prox, proximal. Numbers in parentheses indicate sample sizes (*, n ¼ 6 for TH-IR PSCs in the medial ventral

lip region). For abbreviations, see list.



bright DiI PSCs < HA-IR PSCs (Table 4). The bipolar NOS-

IR PSCs, excluded from the statistical comparison

because they were measured by using a different proce-

dure, had the longest mean process length (Table 4).

Somata diameters were similar across PSC types, except

for NADPHd PSCs, which had significantly smaller somata

(ANOVA, F3,238 ¼ 16.7, P < 0.001, followed by Tukey’s

post hoc comparisons). The bipolar NOS PSCs had similar

somata sizes to the three other bipolar PSCs (Table 4).

DISCUSSION

We provide an unprecedented view of the extent and

complexity of gastropod peripheral sensory neuroanat-

omy in the CSOs of L. stagnalis. We describe both new

PSC types (unipolar putative nitregric PSCs) and greater

detail regarding both morphologies and distributions for

previously described PSCs (bipolar putative catecholami-

nergic, histaminergic, and nitrergic sensory cells). PSCs

were nonuniformly distributed across the surfaces of the

CSOs of Lymnaea stagnalis, with particular concentra-

tions around the mouth (the ventral medial lips), along

the margins of lips and tentacles, the tips of the ten-

tacles, and both a ventral lateral strip and ventral basal

patch on the tentacles (summarized in Fig. 2). Reaching

densities of 5,000 mm�2 in some regions, there were

hundreds to tens of thousands of each of multiple PSC

types per CSO. Thus, the total number of PSCs in all four

CSOs almost certainly dwarfs the �20,000 neurons pres-

ent in the entire CNS (Feng et al., 2009). These results

make it clear that understanding the neural basis of

behavior in L. stagnalis and other gastropods needs to be

extended to include this massive, but relatively ignored

portion of the nervous system.

We identify four types of PSCs. Our backfills extend

previous observations (Nezlin, 1995; van Marle, 1997;

Nakamura et al., 1999b) to three types of bipolar PSCs.

We labeled each population by using at least two addi-

tional methods, and found a fourth population of unipolar

cells also corroborated by two labels. Although we were

unable to achieve satisfactory double labels, several lines

of evidence suggest that the four PSCs types are different

Figure 9. FaGlu fluorescent bipolar putative PSCs in the CSOs of L. stagnalis. A: Fluorescent PSCs had somata (arrowheads) and proc-

esses that penetrated the CSO epithelium (open arrowheads). B: Numerous PSCs were concentrated particularly along the margin of the

lip (arrowheads), and a deeper branching innervation (arrows) was also apparent. C: The medial margin of the tentacles also showed a

high density of fluorescent PSCs, with at least some their processes apparently joining the branching innervation (arrows) of the CSO.

D: The lateral tentacle had few fluorescent PSCs, but did contain an approximately rectilinear pattern of fluorescent fibers. Scale bar ¼20 lm in A,C; 250 lm in B; 50 lm in D.

Wyeth and Croll


populations, including differences in reactivity to labeling

protocols, morphology, ciliation, putative dendritic

lengths, and distributions. Moreover, given the specificity

of the labels (see Materials and Methods), our results

support differences in PSC content that probably corre-

spond to different substances used in neurotransmission.

Putative catecholaminergic PSCsTH-IR PSCs and a matching population of PSCs labeled

with FaGlu were found in both the lips and tentacles.

These cells thus appear to contain catecholamines,

presumably for synaptic transmission, and are apparently

present in all gastropods. TH-IR PSCs have been

Figure 10. Anti-histamine immunoreactive putative bipolar PSCs in the CSOs of L. stagnalis. A: Two HA-IR PSCs, each with one process

that penetrated the epithelium (arrowhead) and another (arrow) that joined the branching innervation of the CSOs. B: The HA-IR PSCs

were concentrated particularly along a ventral strip of lateral tentacle (spanned by the dashed line). C,D: In both the tentacle (C) and lip

(D), the HA-IR PSCs (arrowheads) were not found directly along the margin (not visible at top right in C), but rather inset, along a ventral-

lateral strip in the tentacle and the ventral lateral region of the lip. In both CSOs, the processes of at least some HA-IR PSCs joined the

branching innervation (arrows) of the CSO. In addition, HA-IR fibers without any visible somata were also present (double arrowheads).

Scale bar ¼ 10 lm in A; 400 lm in B; 50 lm in C,D.



observed during development of the lips and tentacles in

L. stagnalis (Voronezhskaya et al., 1999), and we attribute

the reported absence in an earlier study of adult tentacles

(Croll et al., 1999) to poor reagent penetration. Similarly,

in Aplysia spp., an earlier study that found no evidence

for PSCs in CSOs (Salimova et al., 1987), has been

superseded by observations of TH-IR PSCs, corroborated

by histochemistry (Croll, 2001). Moreover, autoradiographic

Figure 11. Anti-nitric oxide synthase immunoreactive and NADPH diaphorase reactive putative PSCs in the CSOs of L. stagnalis. A: Unipo-

lar NOS-IR PSCs (arrowheads) in the tentacle were accompanied by an extensive branching innervation (arrows). Inset: an oblique optical

slice through a confocal stack shows the unipolar cells (arrowheads) and the branching innervation connected to a cluster (c) of bipolar

NOS-IR PSCs; each with a process that penetrated the epithelium (one visible in this plane, p). B: Along the proximal medial margin of the

tentacle, the bipolar NOS-IR PSC somata often occurred in clusters (c), with separated epithelial processes (p) and bundled connections

to branching innervation (arrows). C: NADPHd-R PSCs (arrowheads) were found on both dorsal and ventral surfaces of the tentacle

(both surfaces are visible), along with a deeper NADPHd-R branching innervation (arrows). D: A similar pattern of unipolar PSCs and

innervation was observed in the lip. Inset: the NADPHd-R branching innervation did not appear to connect to the unipolar NADPHd-R

PSCs, whose single process penetrated the epithelium (arrowheads). Scale bar ¼ 200 lm in A, 10 lm in inset to A; 20 lm in B; 100 lmin C,D; 30 lm in inset.

Wyeth and Croll


evidence indicates that some peripheral cells in A. californ-

ica are dopaminergic (Xin et al., 1995). In addition, TH-IR

PSCs were also observed in the CSOs of several other opis-

thobranchs (Croll et al., 2003; Faller et al., 2008), as well as

bivalves (Croll et al., 1997; Smith et al., 1998) and cephalo-

pods (Baratte and Bonnaud, 2009). Thus, the putative cate-

cholaminergic PSCs appear to be widespread in gastropods

and possibly also across the molluscs.

The TH-IR PSCs were broadly distributed in the CSOs,

and were especially dense along the medial margin and tip

of the tentacle, as well as around the lip margin and

mouth. In backfills, PSCs with short epithelial processes

were found in similar locations, and the dimly fluorescent

cells labeled by DiI, although much less dense, were also

similar in morphology. These appear to all be the same

cells, with DiI labeling only a small proportion of the popu-

lation. At least some of the TH-IR PSCs were multiciliated,

as were some of the dim DiI cells and the corresponding

backfilled PSCs. We therefore suggest that the TH-IR PSCs

bear a tuft of motile cilia (because some of the DiI-labeled

cells had beating cilia). We hypothesize that the TH-IR

PSCs project centrally, consistent with the high density of

TH-IR fibers in the CSO nerves (Croll and Chiasson, 1990;

Croll et al., 1999). However, as some PSCs may be intrin-

sic to the CSOs, without complete anti-TH and backfill dou-

ble labels we cannot determine what proportions project

centrally or to other peripheral regions. Interpretation is

further complicated by observation of other peripheral TH-

IR neurons without epithelial processes (Croll et al., 2003;

Faller et al., 2008). Regardless, it appears that multicili-

ated bipolar TH-IR PSCs form a substantial component of

the gastropod peripheral nervous system.

Putative histaminergic PSCsWe provide documentation of HA-IR PSCs in both ten-

tacles and lips of L. stagnalis. Bipolar HA-IR PSCs were

narrowly distributed, concentrated along a ventral lateral

strip of the tentacles and the ventral lateral surface of the

lips. We propose that the HA-IR PSCs correspond to both

the solitary PSCs with long epithelial processes observed

in backfills and bright DiI-labeled PSCs, given the

similarities in size and morphology as well as consistent

distributions (in particular, the ventral lateral strip of the

tentacles). Although the bright DiI cells had significantly

shorter epithelial processes than the HA-IR PSCs, the

differing tissue processing steps for each label could

account for the discrepancy. If these are all the same

cells, then DiI apparently only labels a small subset in the

lips. We suggest that these PSCs bear either a single non-

motile cilium or a cluster of nonmotile cilia, based on their

presence in both DiI and backfill labels. The HA-IR PSCs

probably project centrally, accounting for the high density

of HA-IR in all three CSO nerves (Hegedus et al., 2004)

and their peripheral branches. However, the HA-IR fibers

without somata in the CSOs may also contribute to the

HA-IR fibers in the nerves. Thus, confirmation of projec-

tion patterns for the HA-IR PSCs requires double labels

with backfills. Nonetheless, we suggest that the most

likely scenario is that a population of monociliated bipolar

HA-IR PSCs project directly to the CNS.

Although we were unable to corroborate HA-IR labeling

with a second neurotransmitter-specific method, we expect

that the HA-IR structures contain histamine, given the sup-

porting evidence for specificity of the anti-HA antibody (see

Materials and Methods) and the presence of HA as a neuro-

transmitter in L. stagnalis (Hegedus et al., 2004).

Putative nitrergic PSCsWe observed a widespread population of unipolar

NADPHd-R PSCs, and propose that the similar unipolar

NOS-IR PSCs are the same population of cells. We found

no evidence of ciliation, and thus suggest these are not

the same cells as the nitrergic ciliated epithelial cells

without processes in the CSOs of A. californica (Moroz,

2006). Instead, the epithelial process ended with a

‘‘knob’’-like structure, also noted by Huang and others

(1997) in bipolar NADPHd-R sensory cells in Helix. We

suggest that these unipolar cells may be similar to verte-

brate taste receptors, hair cells, or neuroendocrine cells,

which also lack axons (Roper, 1989; Langley, 1994; Raph-

ael and Altschuler, 2003), with somatic synapses with

other (unidentified) peripheral fibers.

A second population of clustered bipolar NOS-IR PSCs

was also sometimes present. The bipolar NOS-IR PSC

label was particularly ‘‘capricious’’ (Cooke et al., 1994),

which limited our ability to determine their distribution.

Similarly clustered bipolar cells were present in some

backfills in similar locations in the tentacle as well as

other regions in which bipolar NOS-IR PSCs were never

observed. In particular, the dense patch of clustered bipo-

lar PSCs found ventrally on the tentacles (Steffensen

et al., 1993; Nezlin, 1995; van Marle, 1997) has a similar

morphology, yet was never labeled in anti-NOS prepara-

tions. It is possible that there is one type of clustered

PSCs that labeled intermittently with anti-NOS, or, alter-

natively, there may be other clustered PSCs that are

not NOS-IR. No cilia labeled on the bipolar NOS-IR

PSCs, whereas clustered PSCs in backfills were ciliated.

Thus, although we are confident that the bipolar clus-

tered NOS-IR cells are present in at least some areas

of the tentacles, more concrete conclusions regarding

their distribution and ciliation await further observations.

In contrast to our results, bipolar (not unipolar)

NADPHd-R PSCs were described in sectioned lips of

L. stagnalis (Elphick et al., 1995). We propose that sec-

tioning allowed the bipolar NOS-IR PSCs described here



to also label using NADPHd. Consistent with this

explanation are our observations of fainter somata in

whole-mount CNSs with NADPHd-R, when compared

with previous observations of sections (Moroz et al.,

1994b). Alternatively, it is possible that sectioned tis-

sue in the previous study created the impression that

the NADPHd-R branching connected with what were in

fact unipolar NADPHd-R sensory cells. On balance, we

suggest that the evidence supports the presence of

two distinct populations of PSCs, one unipolar, and one

bipolar.

The bipolar NOS-IR PSCs appear to project directly to

the CNS. When bipolar PSCs were present, the NOS-IR

branching innervation clearly connected to the PSCs. In

contrast, both methods consistently labeled fibers that

terminated some distance from the unipolar PSCs. This

might be explained if the bipolar PSCs have distal

portions (including somata and epithelial processes) that

are intermittently labeled with anti-NOS and never with

NADPHd, perhaps in both cases due to inadequate

reagent penetration into CSO margins. Alternatively, a

third set of neurons may have fibers in the CSOs. Either

scenario is consistent with the abundant NADPHd-R and

NOS-IR fibers found in all three CSO nerve trunks of the

CNS (Moroz et al., 1994a; Elphick et al., 1995). Double

labels pairing NADPHd and anti-NOS with each other and

with backfills are required to fully resolve the relationship

between the unipolar and bipolar PSCs and whether the

latter project centrally.

The PSCs labeled with anti-NOS and NADPHd proto-

cols are possibly nitrergic (Moroz, 2000). The unipolar

cells are corroborated by both methods, and the bal-

ance of evidence from NADPHd controls and anti-NOS

immunoreactivity is consistent with the hypothesis that

NOS is present. If they are NOS-containing, then NO is

likely produced by these cells, as direct measurements

of NO and metabolites involved in its production have

been shown in an NADPHd-R cell in the CNS of L. stag-

nalis (Moroz et al., 2005; Patel et al., 2006). We found

no NADPHd-R counterpart to the bipolar NOS-IR PSCs,

but they are corroborated by bipolar NADPHd-R PSCs

in sectioned lip tissues, and neurophysiological assays

further support their nitrergic character (Elphick et al.,

1995). The mismatch in labeling observed here

between NOS immunohistochemistry and NADPHd has

precedent in gastropods, and could result from differ-

ent enzyme isoforms (Moroz et al., 1993; Cooke et al.,

1994; Moroz et al., 1994a; Huang et al., 1997) or prob-

lems with reagent penetration (as noted above). Thus,

although cross-reactivity with another protein remains a

possibility, this population of clustered bipolar PSCs

probably contains NOS and therefore is presumably

nitrergic.

Other cellsReconciling our results with previous PSC descriptions

is complicated by differing labels and observation meth-

ods. Glutamate-containing PSCs have been reported in

sectioned CSOs of L. stagnalis (Hatakeyama et al., 2007),

yet we have been unable to replicate this observation in

either peripheral or CNS whole-mounts (unpublished

observations). FMRFamide-immunoreactive PSCs have

been described in a number of gastropod CSOs (Suzuki

et al., 1997; Wollesen et al., 2007; Faller et al., 2008), but

we were unable to achieve consistent labeling with anti-

FMRFamide (unpublished observations), and thus cannot

confirm whether any are also present in L. stagnalis.

PSCs in the CSOs of L. stagnalis have also been classified

by using electron microscopy (Zylstra, 1972; Zaitseva and

Bocharova, 1981; Roubos and van der Wal-Divendal,

1982), but the majority of cell types have morphologies

that do not match any PSCs described here. Thus, we can

only conclude, supported by our observations of backfills,

that further PSC types are present in the CSOs.

Sensory modalitiesThe morphologies of the cells described here, with

putative dendrites extending through the CSO epithelium,

are consistent with a sensory function. The possible

modalities include mechanosensation, chemosensation

(de Vlieger, 1968; Jager, 1971; Goldschmeding and Jager,

1973; Janse, 1976; Lever, 1977; Zaitseva et al., 1987),

and even photosensation (Chase, 1979; Stoll, 1979). The

bipolar NOS-IR PSCs are probably chemosensory, based

on physiological evidence (Elphick et al., 1995). The uni-

polar NADPHd-R PSCs may also be chemosensory, given

their morphological similarity to putative chemosensory

cells identified in the CSOs of A. californica (Cummins

et al., 2009). In A. californica, the TH-IR PSCs have been

hypothesized to be candidates for mechanosensory cells

mediating low-threshold tactile reflexes (Croll, 2001,

2003). However, here we provide evidence for at least

three other PSC types with unknown function (and which

may or may not be present in A. californica). We see no

obvious way to generate hypotheses for PSC modalities

based on the structures and distributions described here.

Instead, we prefer to emphasize the need for further

physiological experimentation to properly test the sen-

sory nature and modalities of the cells.

Implications and future workWe have mapped a number of possible PSCs in the

CSOs of L. stagnalis, and also matched putative neuro-

transmitters with those cells. The total complement of

PSCs known in L. stagnalis now includes catecholamine-

containing, histamine-containing, glutamate-containing

Wyeth and Croll


(Hatakeyama et al., 2007), and two types of NOS-contain-

ing cells. Further work is now required to verify the

substances these cells use for neurotransmission. None-

theless, meaningful comparisons with other species will

be facilitated by our comprehensive descriptions and

mapping in L. stagnalis. Moreover, the catalog of distribu-

tions, morphologies, and putative neurotransmitters

provided here can help design experiments to assign

modalities to the PSCs. Given the complexity of the pe-

ripheral nervous system described here, we suggest that

these tests are best accomplished with optical recordings

using physiologically sensitive dyes in identifiable PSC

types. The tentacles of L. stagnalis are good first choice

for attempting these experiments, which will be facili-

tated by the distribution and morphological data we pres-

ent for identifying the PSCs. In the long term, the informa-

tion on identified PSC functions will complement our

understanding of central neurons, and help build a more

complete view of neuroethology in the snails and also

provide a basis for comparative work in other gastropods.

ACKNOWLEDGMENTS

We thank P. Panula for a gift of conjugated histamine,

as well as H. Harding, M. Stoyek, S. Whitefield, O. Brau-

bach, C. P. Holmes, the Centre for Medical Digital Imag-

ing at Dalhousie University, and two anonymous

reviewers for assistance.

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