Host Identification and Glochidia Morphology of
Freshwater Mussels from the Altamaha River Basin
Jennifer A. Johnson, Jason M. Wisniewski, Andrea K. Fritts, and Robert B. Bringolf
Southeastern Naturalist, Volume 11, Issue 4 (2012): 733–746
Full-text pdf (Accessible only to subscribers. To subscribe click here.)
2012 SOUTHEASTERN NATURALIST 11(4):733–746
Host Identification and Glochidia Morphology of
Freshwater Mussels from the Altamaha River Basin
Jennifer A. Johnson1, Jason M. Wisniewski2, Andrea K. Fritts1,
and Robert B. Bringolf
Abstract - Recovery of imperiled freshwater mussels requires knowledge of suitable host
fishes and other early life-history traits. We provide quantitative host information for 6
mussel species from the Altamaha River Basin, GA, 3 of which previously had no host information.
Glochidia of Alasmidonta arcula (Altamaha Arcmussel) metamorphosed on 2
species of suckers (Moxostoma spp.); Elliptio hopetonensis (Altamaha Slabshell) on Lepomis
macrochirus (Bluegill), Pimephales promelas (Fathead Minnow), and Micropterus
salmoides (Largemouth Bass); E. shepardiana (Altamaha Lance) on 2 species of Bullheads
(Ameiurus spp.) and L. macrochirus; Lampsilis dolabraeformis (Altamaha Pocketbook)
on Bluegill and Largemouth Bass; and L. splendida (Rayed Pink Fatmucket) and Villosa
delumbis (Eastern Creekshell) on Largemouth Bass. We also provide descriptions of glochidia
morphology for the above mussel species and E. spinosa (Altamaha Spinymussel).
Glochidia were correctly identified to species in 88.7% of cases by discriminant function
analysis of 3 shell dimensions. Glochidia morphology may be useful for identification of
glochidia attached to wild fish, thereby providing additional host information.
Freshwater mussels (family Unionidae) provide critical ecosystem services
and often dominate the benthic biomass in minimally impacted streams (Strayer
et al. 2004). Live mussels and empty shells provide habitat for other invertebrates
and fish, and as filter feeders, they influence nutrient cycling by linking the water
column and the substrate (Spooner and Vaughn 2006). Nearly 300 species of
unionids occur in North America, but unfortunately they are highly imperiled,
with approximately 70% of North American species being considered of conservation
concern (Neves et al. 1997, Williams et al. 1993). Factors thought to
contribute to the decline of mussels include sedimentation, pollution, urbanization,
and habitat fragmentation (Williams et al. 1993). Because nearly all mussel
larvae (glochidia) are obligate parasites on fish, declines in host fish populations
may also contribute to mussel declines.
The Altamaha River in Georgia (drainage area = 36,976 km2) is inhabited by
approximately 18 mussel species, of which 7 are endemic (Johnson 1970). At
least 3 of the endemic species, Alasmidonta arcula Lea (Altamaha Arcmussel),
Pyganodon gibbosa Say (Inflated Floater), and Elliptio spinosa Lea (Altamaha
Spinymussel), are declining (Dinkins et al. 2004, Keferl 1981, O’Brien 2002a,
Skelton et al. 2002, Wisniewski et al. 2005), and E. spinosa was listed as
1Warnell School of Forestry and Natural Resources, University of Georgia, Athens, GA
30602. 2Nongame Conservation Section, Wildlife Resources Division, Georgia Department
of Natural Resources, Social Circle, GA 30025. *Corresponding author - bringolf@
734 Southeastern Naturalist Vol. 11, No. 4
endangered in 2011 under the US Endangered Species Act. Fish hosts of mussel
species in the Altamaha River are poorly known, and of the 3 declining endemic
species, limited host information is available only for A. arcula. Identification of
host fishes allows managers to determine if appropriate host species are present
in the river, and knowledge of suitable hosts is also necessary for captive propagation
of mussels, which can produce juveniles for population augmentation or
toxicity testing. The Georgia State Wildlife Action Plan (2005; http://www.georgiawildlife.
com/conservation/wildlife-action-plan) identified knowledge of early
life histories of mussels endemic to the Altamaha River Basin as a high priority
need for recovery of these species. Description of glochidia morphology also is
necessary to identify patterns of glochidia occurrence on wild fishes, to inform
phylogenetic relationships among species, and to provide additional characters
for identification. In this study, we identify host fish and describe glochidia
morphology of 7 mussels from the Altamaha River Basin: A. arcula, E. spinosa,
E. hopetonensis Lea (Altamaha Slabshell), E. shepardiana Lea (Altamaha
Lance), Lampsilis dolabraeformis Lea (Altamha Pocketbook), L. splendida Lea
(Rayed Pink Fatmucket), and Villosa delumbis Conrad (Eastern Creekshell).
Gravid female mussels were collected from the mainstem Altamaha River in
2008–2009 (Table 1) by visual and tactile searches with SCUBA and snorkel. All
species except A. arcula and E. spinosa were collected downstream of Oglethorpe
Bluff Landing, ≈12 km north of Jesup, GA (Fig. 1); A. arcula and E. spinosa
were collected upstream of Upper Wayne Landing, ≈17.1 km SSW of Glennville,
GA. Mussels were gently pried open to detect marsupial swelling, which indicate
females are gravid and brooding glochidia. On each sample date, at least 10
individuals of each species were examined, and if no gravid mussels were found,
inspections ceased. Gravid females of each species were transported in coolers
to the University of Georgia. To minimize premature release of glochidia, mussels
were maintained in dechlorinated tap water at 10–12 ºC in a Living Stream
(Frigid Units, Inc., Toledo, OH). Mussels were fed weekly with a mixture of concentrated
microalgae (Reed Mariculture Instant Algae® Shellfish Diet and Nanno
Table 1. Collection dates and water temperature at time of collection of gravid female mussels from
the Altamaha River.
Species Number collected Date collected Water temperature (oC)
Alasmidonta arcula 5 12 Nov 2009 16
Elliptio hopetonensis 5 18 June 2009 29
Elliptio shepardiana 5 18 June 2009 29
Elliptio spinosaA 1 18 May 2009 21
Lampsilis dolabraeformis 5 18 June 2009 29
Lampsilis splendida 2 1 Oct 2008 26
Villosa delumbis 2 1 Oct 2008 26
AGlochidia immature at time of collection; mature glochidia released on 10 June 2009.
2012 J.A. Johnson, J.M. Wisniewski, A.K. Fritts, and R.B. Bringolf 735
3600 Nannochloropsis). Glochidia were extracted from gravid females within 2
months for morphological characterization and host trials.
Fish for host trials were obtained from state, federal, and private fish hatcheries
in the southeastern US, or by electrofishing or seining in ponds and streams in
the Altamaha basin where mussels do not occur. Small (presumably young) individuals
of each fish species were used when possible to maximize likelihood of
successful metamorphosis by glochidia (Strayer 2008). Based on length at capture,
all Micropterus salmoides (Largemouth Bass) and Lepomis macrochirus (Bluegill)
used were assumed to be young of year. Ideally, fish used in host trials have not
had any previous exposure to mussels because fish develop acquired immunity to
glochidia following exposure (Dodd et al. 2005, 2006; Meyers et al. 1980; Zale and
Neves 1982). Fish species used in each trial depended on availability and were not
the same for each mussel species (Tables 2, 3).
Glochidia were obtained from 1 to 4 females of each species by gently
flushing the marsupial gills with water from a 5-ml syringe. Elliptio spinosa,
E. hopetonensis, and E. shepardiana released mature glochidia within one month
of transport to the lab, so glochidia of these species were collected without the
use of a syringe. Subsamples of 50–100 glochidia per female were tested for
viability by exposure to a saturated sodium chloride solution; viable glochidia
quickly closed their valves upon exposure to the solution. Only female mussels
with glochidia viability of >90% were used for host trials. Potential host fishes
Figure 1. Altamaha River Basin,
GA and approximate location
(oval) of freshwater
mussel collections from the
mainstem Altamaha River.
736 Southeastern Naturalist Vol. 11, No. 4
were separated by species and exposed to glochidia for one hour in aerated 19-L
buckets. Each bucket contained 1–22 individuals of each fish species (Tables
2, 3), and a target glochidia concentration of 4000/L. For E. spinosa, a limited
amount of viable glochidia were available (from a single female), so glochidia
were pipetted directly on the fish gills to increase the probabi lity of attachment.
Glochidia-infested fish were removed from the glochidia suspension, gently
rinsed to remove any unattached glochidia, and placed in holding (monitoring)
tanks in a recirculating aquaculture system (AHAB; Aquatic Habitats Inc., Apopka,
FL). The AHAB unit is an array of self-cleaning tanks through which water
flows and then re-circulates back into a main sump for treatment with activated
carbon and UV sterilization. Whenever possible, we placed one fish per tank, but
unfortunately this was not feasible for all species because of limited space; no
tanks contained more than 3 fish (always the same species). Water temperatures
ranged from 21–25 ºC during all host trials. Filter cups (5-cm PVC pipe with
153-μm-mesh screen on one end) were fitted to the outlet of each tank to capture
dead glochidia and metamorphosed juveniles as they were sloughed from the
Table 2. Summary of Alasmidonta arcula host trial. Metamorphosis success is reported as mean
± 95% confidence interval. Juv = juveniles produced, Period = period in days of juvenile release,
# = number of fish used.
Fish species success (%) Juv Period #
Moxostoma robustum (Cope) (Robust Redhorse) 4.6 ± 0.63 61 8–13 5
Moxostoma rupiscartes Jordan and Jenkins (Striped Jumprock) 0.12 ± 0.11 4 10–11 7
Acipenser brevirostrum Lesueur (Shortnose Sturgeon) 0 0 - 2
Acipenser fulvescens Rafinesque (Lake Sturgeon) 0 0 - 5
Acipenser oxyrinchus Mitchill (Atlantic Sturgeon) 0 0 - 2
Ameiurus brunneus (Jordan) (Snail Bullhead) 0 0 - 2
Ameiurus natalis (Leseur) (Yellow Bullhead) 0 0 - 1
Cyprinus carpio L. (Common Carp) 0 0 - 2
Etheostoma inscriptum (Jordan and Brayton) (Turquoise Darter) 0 0 - 3
Hypentelium nigricans (Lesueur) (Northern Hogsucker) 0 0 - 6
Ictalurus punctatus (Rafinesque) (Channel Catfish) 0 0 - 5
Lepomis auritus (L.) (Redbreast Sunfish) 0 0 - 1
Lepomis cyanellus Rafinesque (Green Sunfish) 0 0 - 5
Lepomis macrochirus Rafinesque (Bluegill Sunfish) 0 0 - 7
Lepisosteus osseus (L.) (Long Nose Gar) 0 0 - 3
Minytrema melanops (Rafinesque) (Spotted Sucker) 0 0 - 3
Micropterus salmoides (Lacépède) (Largemouth Bass) 0 0 - 3
Morone chrysops (Rafinesque) (White Bass) 0 0 - 5
Moxostoma collapsum (Cope) (Notch-Lip Sucker) 0 0 - 1
Nocomis leptocephalus (Girard) (Bluehead Chub) 0 0 - 4
Notemigonus crysoleucas (Mitchill) (Golden Shiner) 0 0 - 3
Notropis hudsonius (Clinton) (Spottail Shiner) 0 0 - 1
Notropis lutipinnis (D.S. Jordan & Brayton) (Yellowfin Shiner) 0 0 - 5
Notropis rubescens Bailey (Rosyface Chub) 0 0 - 4
Noturus leptacanthus D.S. Jordan (Speckled Madtom) 0 0 - 1
Pimephales promelas Rafinesque (Fathead Minnow) 0 0 - 4
Plyodictis olivaris (Rafinesque) (Flathead Catfish) 0 0 - 2
Semotilus atromaculatus (Mitchill) (Creek Chub) 0 0 - 4
2012 J.A. Johnson, J.M. Wisniewski, A.K. Fritts, and R.B. Bringolf 737
fish. Beginning one day after exposure (day 1), filter cups were checked daily for
7 days, and dead glochidia and metamorphosed juveniles were counted and photographed.
After day 7, the cups were checked every other day. Filtered material
Table 3. Summary of host trials by species of mussel from the Altamaha River, GA. Metamorphosis
success is reported as mean ± 95% confidence interval. Period = period in days of juvenile
release, # = number of fish used.
Mussel species/ Metamorphosis Juveniles
fish species Common name success (%) produced Period #
Lepomis macrochirus Bluegill 3.7 ± 1.0 49 7–8 8
Pimephales promelas Fathead Minnow 3.1 ± 0.7 55 7 4
Micropterus salmoides Largemouth Bass 0.8 ± 0.4 16 7 4
Acipenser fulvenscens Lake Sturgeon 0 0 - 3
Cyprinus carpio Common Carp 0 0 - 4
Ictalurus punctatus Channel Catfish 0 0 - 3
Ameiurus nebulosus Brown Bullhead 45.2 ± 35.8 378 11–17 2
Ameiurus natalis Yellow Bullhead 18.9 17 11–14 1
Lepomis macrochirus Bluegill 2.2 ± 1.4 4 11–12 6
Moxostoma robustum Robust Redhorse 0.1 ± 0.06 1 17 3
Acipenser fulvenscens Lake Sturgeon 0 0 - 2
Hypentelium nigricans Northern Hogsucker 0 0 - 2
Nocomis leptocephalus Bluehead Chub 0 0 - 4
Lepomis microlophus Redear Sunfish 0 0 - 5
Pimephales promelas Fathead Minnow 0 0 - 5
Notropis hudsonius Spottail Shiner 0 0 - 1
Notemigonus crysoleucas Golden Shiner 0 0 - 1
Notropis lutipinnis Yellowfin Shiner 0 0 - 15
Acipenser fulvenscens Lake Sturgeon 0 0 - 4
Cyprinus carpio Common Carp 0 0 - 4
Ictalurus punctatus Channel Catfish 0 0 - 4
Lepomis auritus Redbreast Sunfish 0 0 - 4
Lepomis macrochirus Bluegill 0 0 - 3
Micropterus salmoides Largemouth Bass 0 0 - 4
Minytrema melanops Spotted Sucker 0 0 - 4
Morone chrysops White Bass 0 0 - 1
Morone saxatilis Striped Bass 0 0 - 4
Pimephales promelas Fathead Minnow 0 0 - 2
Micropterus salmoides Largemouth Bass 74.8 ± 3.6 1209 19–29 4
Lepomis macrochirus Bluegill 1.5 ± 0.1 23 10–19 22
Acipenser fulvenscens Lake Sturgeon 0 0 - 3
Ictalurus punctatus Channel Catfish 0 0 - 2
Notemigonus crysoleucas Golden Shiner 0 0 - 5
Pimephales promelas Fathead Minnow 0 0 - 4
Micropterus salmoides Largemouth Bass 43.0 ± 5.5 2352 13–24 5
Micropterus salmoides Largemouth Bass 73.1 ± 1.5 4673 12–24 18
738 Southeastern Naturalist Vol. 11, No. 4
was gently rinsed into a Bogorov tray and examined under a stereomicroscope.
Juveniles were identified by the presence of tissues such as gills, foot, and heart.
To determine if juveniles were alive, we observed them for foot movement, heartbeat,
or valve closure. We checked filter cups every 2 days until no glochidia or
juveniles were observed for 5 consecutive days. When fish mortality occurred,
deceased fish were examined for glochidia. No high infestations were observed
on dead fish, and data from these fish was not included in the final analysis of
We quantified juvenile metamorphosis (%M) for individual fish as ([number
of juveniles/ (juveniles + glochidia)] x 100). When more than one individual of
a fish species was used per tank, we determined the total %M for the tank. We
then calculated the mean %M across replicates (tanks) for each species. Initial
glochidia attachment rates were determined by summing the total number of
sloughed glochidia and juveniles that were recovered from each tank.
A sub-sample (n = 100–150) of glochidia from each individual mussel was
fixed in formalin for at least 24 hours and then stored in 95% ethanol. Morphological
measurements were made for 25 glochidia/female for each species
(Table 4). Glochidia were photographed at magnifications of 16–50X with a stereomicroscope
(Leica MZ 7.5, Leica Microsystems, Wetzlar, Germany) equipped
with a digital camera (Leica DCF 290, Leica Microsystems, Wetzlar, Germany).
Glochidia shape classifications were based on previous descriptions (Hoggarth
1999, Hornbach et al. 2010). Glochidia length (parallel to hinge), height (perpendicular
to hinge), and hinge length (Hoggarth 1999, Kennedey and Haag 2005)
were measured with image analysis software (Leica LAS, Leica Microsystems,
Wetzlar, Germany). Differences in mean length, height, and hinge-length measurements
were compared among species with 3 separate analysis of variance
(ANOVAs) followed by Tukey’s test to identify differences between species for
each measurement (α = 0.05). We also examined the utility of shell measurements
to identify glochidia by species with a discriminant function analysis (DFA) as
previously described by Kennedy and Haag (2005). Briefly, all measurements
were transformed (log10[x + 1]) to achieve normality, and we derived quadratic
discriminant functions for each species because variance-covariance matrices
Table 4. Glochidia (n = 25 per female) measurements (mean ± 95% confidence interval) for
freshwater mussels of the Altamaha River, GA. Within a column (shell dimension), different superscripted
capital letters indicate significant differences among species (Tukey’s test, α = 0.05);
species with the same letter within a column were not significan tly different.
Species # of females Height (μm) Length (μm) Hinge (μm)
Alasmidonta arcula 4 A309 ± 14.3 A274 ± 5.4 A212 ± 4.1
Elliptio hopetonensis 1 B199 ± 4.7 B186 ± 3.8 B134 ± 2.4
Elliptio shepardiana 4 C259 ± 6.5 C217 ± 4.0 C153 ± 3.1
Elliptio spinosa 1 B208 ± 3.8 D197 ± 3.0 B133 ± 3.7
Lampsilis dolabraeformis 4 D239 ± 5.2 E207 ± 3.3 D100 ± 3.4
Lampsilis splendida 4 C268 ± 3.0 C222 ± 4.0 D97 ± 4.2
2012 J.A. Johnson, J.M. Wisniewski, A.K. Fritts, and R.B. Bringolf 739
were unequal (c2 = 92.7, df = 30, P < 0.0001; Morrison 1976). We used cross-validation
scores to determine identification success for each individual glochidium
and reported results as the % of total number of measured glochidia identified
correctly. All statistical analyses were performed with SAS (version 8.2, SAS
Institute, Cary, NC).
Glochidia ultrastructure (i.e., presence of microstylets, interior/exterior valve
sculpture) was described for L. dolabraeformis, E. shepardiana, E. spinosa, and
A. arcula from scanning electron microscope images (SEM, Zeiss 1450EP, Zeiss
SMT, Peabody, MA). We were unable to describe ultrastructure for glochidia
of L. splendida and V. delumbis due to the low quality of preserved specimens.
Glochidia samples were mounted on SEM stubs with carbon adhesive tabs (EMS,
Hatfield, PA), and a SPI Module Sputter Coater (SPI Supplies, Inc. West Chester,
PA) was used to coat samples with 20 μm of gold. Specimens were then examined
under the SEM run at 20 Kev with a probe size of 450 uA.
Juvenile A. arcula (61 individuals total, %M = 4.6) were produced by all 4
individuals of Moxostoma robustum Cope (Robust Redhorse) and 4 A. arcula
juveniles (%M = 0.8) were produced from 1 of the 7 Moxostoma rupiscartes
(Striped Jumprock). No juvenile A. arcula were produced from 26 other fish
species tested (Table 2). Juvenile E. hopetonensis were produced from Bluegill
(%M = 3.7), Pimephales promelas (Fathead Minnow; %M = 3.1), and Largemouth
Bass (%M = 0.8), but 3 other species were non-hosts (Table 3). Juvenile
E. shepardiana were produced by Ameiurus nebulosus Lesueur (Brown Bullhead;
%M = 45.2), A. natalis (Yellow Bullhead; %M = 18.9) and Bluegill
(%M = 2.2%). A single E. shepardiana juvenile was produced from a Robust
Redhorse (%M = <0.1%), and no juveniles were produced from 8 other fish
species (Table 3). Juvenile L. dolabraeformis were produced from Largemouth
Bass (%M= 74.8%) and Bluegill (%M = 1.5), but no juveniles were produced
from 4 additional fish species (Table 3). Largemouth Bass also produced juvenile
L. splendida (% M = 43) and V. delumbis (%M = 73.1); no other fish
species were tested with L. splendida and V. delumbis (Table 3). None of the 10
fish species tested produced juvenile E. spinosa (Table 3). Eight fish species
sloughed 100% of the attached E. spinosa glochidia within 3 days after initial
glochidia exposure (data not shown), but 4 E. spinosa glochidia remained attached
on Acipenser fulvescens (Lake Sturgeon) and 5 on Lepomis auritus
(Redbreast Sunfish) until 5 days after attachment.
Glochidia morphology and shell ultrastructure
Glochidia height (F5,149 = 161.3, P < 0.0001), length (F5,149 = 323.5, P < 0.0001),
and hinge length (F5,149 = 530.4, P < 0.0001) all varied significantly among the 6
species (Table 4). The 95% confidence interval for a given shell dimension overlapped
with 0–2 other taxa (Table 4). The DFA correctly classified 133 (88.7%)
of the 150 glochidia in the data set, and correct classification percentages by
740 Southeastern Naturalist Vol. 11, No. 4
species ranged from 60% to 100% (Table 5). Correct classification was 100%
for 3 of the 6 taxa. Glochidia were misclassified as 0–2 other taxa, and the most
common misclassification was E. hopetonensis as E. spinosa (40%). Conversely,
E. spinosa were misclassified as E. hopetonensis in 5 of 25 cases (20%). All other
misclassifications were ≤4%.
Glochidia shell structure varied by species. Alasmidonta arcula glochidia
were pyriform and contained a styliform hook ventrally located on each pitted
valve (Fig. 2). Lampsilis dolabraeformis had a pitted subelliptical valve with
a ventral edge (flange) covered with micropoints (Fig. 3). Lampsilis splendida
glochidia were also subelliptical, but we were unable to determine ultrastructure
Table 5. Identification success (%) for glochidia of 6 mussel taxa from the Altamaha River Basin.
Values were determined with cross-validated scores of quadratic discriminant functions for 25
glochidia of each species. Numbers in parenthesis are % of glochidia misclassified for the given
Species Correct (%) Misclassified as
Alasmidonta arcula 100 -
Elliptio hopetonensis 60 E. spinosa (40)
Elliptio shepardiana 100 -
Elliptio spinosa 76 E. hopetonensis (20), L. dolabraeformis (4)
Lampsilis dolabraeformis 100 -
Lampsilis splendida 96 L. dolabraeformis (4)
Figure 2. Scanning electron micrographs of Alasmidonta arcula glochidia A) exterior,
200X, B) interior 200X, C) side view, 160X, and D) detail of styliform hook, 1100X.
Scale bars = 100μm.
2012 J.A. Johnson, J.M. Wisniewski, A.K. Fritts, and R.B. Bringolf 741
with SEM. Glochidia of E. hopetonensis had a depressed subelliptical shape.
Elliptio shepardiana glochidia had a pitted valve structure and a depressed subelliptical
shape with a ventral flange extended beyond the gill margin covered
with numerous microstylets (Fig. 4A). Similar to the other members of the genus
Elliptio, E. spinosa was also pitted with a depressed subelliptical shape and the
ventral flange was covered with microstylets, but not extended (Fig. 4B). We
also observed several subcylindrical white conglutinate packets (with immature
glochidia) released by E. spinosa in the lab; conglutinate length was 18 mm and
width was 5 mm.
We produced quantitative host information for 6 mussel species from the Altamaha
River Basin, GA, 3 of which previously had no host information. Our finding
that robust Redhorse may serve as a host for A. arcula is noteworthy because we
are aware of only one other report of a mussel-host fish relationship involving 2
imperiled species (Fritts et al. 2012). Robust Redhorse were historically abundant
in the Altamaha Basin but are now listed as state endangered. They were thought
to be extinct for 122 years until their “rediscovery” in 1991 in the Altamaha Basin
and subsequently in other Atlantic Slope drainages north to Virginia (Grabowski
and Jennings 2009). The relatively low metamorphosis rate (4.6%) of A. arcula
on Robust Redhorse may not be indicative of the importance of this fish species as
Figure. 4. Scanning electron micrographs of the exterior valves of A) Elliptio shepardiana,
200X, and B) Elliptio spinosa, 200X. Scale bars = 100 μm.
Figure. 3. Scanning electron micrographs of Lampsilis dolabraeformis glochidia A) interior
flange, 1180X, B) interior, 200X, and C) valve exterior, 200X. The wavy margin
(dissociation of pellicle from shell) is an artifact of preservation.
742 Southeastern Naturalist Vol. 11, No. 4
a host because host trials were performed at warmer temperatures than likely experienced
by glochidia and fish in the river during the time of natural encystment.
Gravid A. arcula were collected in November when water temperature was 16 oC;
thus, under natural conditions glochidia would develop on host fish when water
temperatures were likely at or below this temperature. Unfortunately, we lacked
the ability to chill the water in our trial, and temperatures were 22–24 oC. The warm
temperatures may have created sub-optimal conditions for glochidia, resulting
in lower %M. The only other species that produced any A. arcula juveniles was
Striped Jumprock, suggesting that A. arcula may be a host specialist, using members
of the sucker family (Catastomidae). Other suckers did not produce juveniles
in our trials, but these tests should be repeated at cooler water temperatures that
more closely mimic natural conditions before any definitive conclusions are made.
Further, we have tested only a fraction of the 93 extant fish species in the Altamaha
River system, and other fishes may be important hosts. Other investigators have
previously reported that A. arcula metamorphose on Gambusia holbrooki Girard
(Eastern Mosquitofish; O’Brien 2002b). Similar to other mussel species, identification
of host(s) for A. arcula is imperative for successful conservation; therefore,
efforts to identify additional hosts should continue.
O’Brien (2002b) reported the only other published host information for Altamaha
mussel species. In that study, juvenile E. hopetonensis were produced
from Eastern Mosquitofish and Bluegill, but not from Largemouth Bass. In the
present study, juvenile E. hopetonensis were produced by Bluegill, Largemouth
Bass, and Fathead Minnows. O’Brien (2002b) reported that L. dolabraeformis
juveniles were produced from Eastern Mosquitofish and Largemouth Bass, but
not from Bluegill. In the present study, juvenile L. dolabraeformis were produced
on Largemouth Bass and Bluegill to a lesser extent.
One of the objectives for this study was to identify a host fish for the federally
endangered E. spinosa. Unfortunately, suitable hosts for E. spinosa remain
unknown at this time. The major limiting factor in our trials was the inability
to collect gravid females. In 2009 and 2010, record high flows in the Altamaha
limited search efforts in April–June, the period when gravid E. spinosa had been
collected by other investigators (P. Johnson, Alabama Department of Conservation
and Natural Resources Alabama Aquatic Biodiversity Center, Marion, AL, pers.
comm.). Future host trials with E. spinosa will depend upon a source of gravid females.
One option is to collect E. spinosa throughout the year and relocate them to
a centralized location where the chances of recapture are greater. Another option
is to attach external sonic tags or passive integrated transponder (PIT) tags to the
mussels to enhance the chances of recovery during periods of gravidity. Additionally,
E. spinosa may be collected throughout the year and returned to a culture
facility to determine if they will undergo fertilization and brooding in captivity.
A number of factors can influence metamorphosis success in host trials, including
those in the present study. Mussels may have higher metamorphosis rates
on fish with which they co-occur than fish of the same species from other basins
(Strayer 2008). Metamorphosis success also appears to be higher on smaller and
younger fish than older and larger fish of the same species even if the larger fish
2012 J.A. Johnson, J.M. Wisniewski, A.K. Fritts, and R.B. Bringolf 743
have never had prior glochidia exposure (Dodd et al. 2006). Further, metamorphosis
rates may be lower in wild fish collected from streams where mussels
occur because fish may develop acquired immunity to glochidia (Dodd et al.
2005). In the present study, not all fish species we tested occur in the Altamaha
basin (e.g., Lake Sturgeon), or were native to the basin (e.g., Fathead Minnow),
but we attempted to maximize the number of species tested for potential propagation
of imperiled species. For studies with the exclusive goal of identification of
natural hosts, we recommend collecting small (young) fish that co-occur in the
basin where mussels are collected.
Host studies frequently report only numbers of juveniles produced per fish
species, not the metamorphosis rate on an individual fish or for a fish species.
Numbers of juveniles produced provides a qualitative assessment of hosts, but
evaluation of %M is essential for quantitative assessment and defensible classification
(e.g., “primary”, “secondary”, etc.) of suitability of host species. We
recommend that future host studies report %M for each fish species, and for individual
fish when possible, to provide a quantitative assessment of the relative
potential contribution for metamorphosis on a particular species of fish. Reporting
of infestation rates provides additional valuable information that, along with
%M, would provide a holistic view of relative host suitability .
We provided some of the first detailed descriptions for glochidia of mussels
from the Altamaha River basin. Shell measurements were similar among some
species we examined, but no species were similar in all 3 dimensions. A qualitative
assessment of shell measures among species, comparison of overlapping 95% confidence
intervals, was generally useful for species identification; overlap of height
and length among species was minimal. The DFA was successful for distinguishing
the correct species in >88% of the cases. The most common error (10 of 17 total)
was E. hopetonensis misidentified as E. spinosa. Elliptio shepardiana, the only
other member of the genus Elliptio in the present study, was larger in all 3 shell dimensions
than the other 2 members of the genus and was readily distinguished by
DFA from all other taxa (100% correct identification). Lampsilis dolabraeformis
and L. splendida were correctly identified in 98% (49 of 50) of cases and were distinguished
by the shortest hinge lengths of all taxa examined. With DFA, glochidia
from all 6 of the taxa tested here could be placed in groups of ≤2 taxa. Thus, consistent
with previous findings (Kennedy and Haag 2005), glochidia morphology may
be useful for identifying glochidia, such as those attached to wild fish. However,
some glochidia may remain indistinguishable by analysis of morphometric data,
and use of molecular techniques such as DNA barcoding may allow positive identification
(Boyer et al. 2011). Although molecular technology is rapidly evolving,
analysis of morphometrics is currently more readily accessible and economical
than genetic approaches. Further efforts should be made to describe glochidia
morphology of the other species in the Altamaha Basin, as morphology may also
be a potential method to examine problematic taxonomic relationships of mussels
within the Altamaha Basin and other basins in the South Atlantic Slope. We also
recommend that future studies seek to more completely describe the variability in
glochidia shell measures from individual gravid females of a given species.
744 Southeastern Naturalist Vol. 11, No. 4
Dimensions and features of A. arcula glochidia were consistent with other
members of the genus Alasmidonta (Ortmann 1911). Species in this genus exhibit a
styliform hooked shell and tend be larger in size than hookless glochidia (Barnhart
et al. 2008, Williams et al. 2008). Hooked glochidia often attach to fins and body
surfaces of hosts whereas hookless glochidia generally attach to gills (Barnhart et
al. 2008). Measurements and morphology of E. hopetonensis and E. shepardiana
were similar to those previously reported for these species by O’Brien et al. (2003),
while morphology of viable E. spinosa glochidia had not been reported prior to this
study. The size and shape of E. spinosa are generally similar to those previously
reported for other members of the genus Elliptio (O’Brien et al. 2003, Williams
et al. 2008); however, the ventral flanges of E. spinonsa and E. shepardiana were
distinctly different in length, which may be helpful when trying to distinguish
between these species. Many physical characteristics of L. dolabraeformis and
L. splendida glochidia were similar to other species in the genus Lampsilis. For
example, L. dolabraeformis and L. splendida were comparable to Hamiota subangulata
Lea (Shinyrayed Pocketbook) glochidia, which had an average height of
261 ± 7 μm (O’Brien and Brim-Box 1999). However, L. straminea Conrad (Rough
Fatmucket), L. ornata Conrad (Southern Pocketbook), and L. teres Rafinesque
(Yellow Sandshell) all were markedly smaller than both L. dolabraeformis and
L. splendida (Kennedy and Haag 2005). When feasible, future analyses of glochidia
morphology should be conducted using a scanning electron microscope, which
can provide precise measurements and detailed images of shell ultrastructure that
are not feasible with traditional light microscopy techniques.
Additional studies on the reproductive biology and early life history of
declining species in the Altamaha River are warranted. Efforts to protect imperiled
species will be greatly enhanced by knowledge of spawning and brooding
periods, optimal brooding temperatures, host fish, and descriptions of glochidia
morphology. Knowledge of mussel early life history may provide insight into
causes of specific mussel population declines (e.g., loss of fish host or preferred
habitat) and may be used for development of propagation programs for population
augmentation or reintroduction for restoration and preservation of freshwater
Funding for this project was provided by the Georgia Department of Natural Resources,
Wildlife Resources Division Nongame Conservation Section. Additional funds were
provided by the Altamaha River Cooperative and the United States Fish and Wildlife
Service. We are indebted to many people who provided assistance in the laboratory and
field including Dr. Chris Barnhart, Kaitlin Brotman, Mieko Camp, Derek Colbert, Julie
Creamer, Peter Hazelton, Jimmy Rickard, Colin Shea, Amos Tuck, and Deb Weiler. Bob
Ratajczak provided valuable assistance with statistical analyses in addition to laboratory
support. Dr. John Shields in the University of Georgia Center for Advanced Ultrastructural
Research provided invaluable assistance with scanning electron microscopy. We
are also grateful to Brian Simmons and the staff at Owen and Williams Fish Farm for
donating Largemouth Bass for host trials. We thank two anonymous reviewers for their
constructive comments to improve the manuscript.
2012 J.A. Johnson, J.M. Wisniewski, A.K. Fritts, and R.B. Bringolf 745
Barnhart, M., W. Haag, and W. Roston. 2008. Adaptations to host infection and larval
parasitism in Unionoida. Journal of the North American Benthological Society
Boyer, S.L., A.A. Howe, N.W. Juergens, and M.C. Hove. 2011. A DNA-barcoding approach
to identifying juvenile freshwater mussels (Bivalvia: Unionidae) recovered
from naturally infested fishes. Freshwater Science 30(1):182–194 .
Dinkins, G.R., J.R. Dinkins, J.E. Daniel. 2004. Survey for native mussels with a focus
on Altamaha Spinymussel (Elliptio spinosa) and Altamaha Arc-mussel (Alasmidonta
arcula) in approximately 15 km of lower Ocmulgee River, Coffee/Telfair/Jeff Davis
counties, Georgia. The Nature Conservancy, Darien, GA.
Dodd, B., M.C. Barnhart, C.L. Rogers-Lowery, T. Fobian, and R. Dimock. 2005. Crossresistance
of Largemouth Bass to glochidia of unionid mussels. Journal of Parasitology
Dodd, B., M. Barnhart, C. Rogers-Lowery, T. Fobian, and R. Dimock. 2006. Persistence
of host response against glochidia larvae in Micropterus salmoides. Fish and Shellfish
Fritts, A.K., M.W. Fritts II, D.L. Peterson, D.A. Fox, and R.B. Bringolf. 2012. Critical
linkage of imperiled species: Gulf Sturgeon as host for Purple Bankclimber mussels.
Freshwater Science 31(4):122–1232.
Grabowski, T.B., and C.A. Jennings. 2009. Post-release movements and habitat use of
Robust Redhorse transplanted to the Ocmulgee River, Georgia. Aquatic Conservation-
Marine and Freshwater Ecosystems 19(2):170–177.
Hoggarth, M.A. 1999. Descriptions of some of the glochidia of the unionidae (Mollusca
: Bivalvia). Malacologia 41(1):1–118.
Hornbach, D., V.J. Kurth, and M.C. Hove. 2010. Variation in freshwater mussel
shell sculpture and shape along a river gradient. American Midland Naturalist
Johnson, R.I. 1970. The systematics and zoogeography of the Unionidae (Mollusca: Bivalvia)
of the southern Atlantic Slope Region. Bulletin of the Museum of Comparative
Keferl, E.P. 1981. A survey of the naiads of the Ohoopee River, Georgia. The Bulletin of
the American Malacological Union 11–15.
Kennedy, T.B., and W.R. Haag. 2005. Using morphometrics to identify glochidia from a
diverse freshwater mussel community. Journal of the North American Benthological
Meyers, T.R., R.E. Millemann, and C.A. Fustish. 1980. Glochidiosis of salmonio fishes.
IV. Humoral and tissue responses of Coho and Chinook Salmon to experimental infection
with Margaritifera margaritifera (L.) (Pelecypoda: Margaritanidae). The Journal
of Parasitology 66(2):274–281.
Morrison, D.F. 1976. Multivariate Statistical Methods. McGraw Hill, New York, NY.
Neves, R., A. Bogan, J. Williams, S. Ahlstedt, and P. Hartfield. 1997. Status of aquatic
mollusks in the southeastern United States: A downward spiral of diversity. Southeast
Aquatic Research Institute, Aquatic Fauna in Peril: The Southeastern Perspective,
Special Publication 1:43–85.
O’Brien, C. 2002a. A survey of the Altamaha Spinymussel (Elliptio spinosa) and Altamaha
Arcmussel (Alasmidonta arcula) in the Altamaha and Ocmulgee Rivers, Georgia.
Final Report to the Georgia Department of Natural Resources, Social Circle, GA.
746 Southeastern Naturalist Vol. 11, No. 4
O’Brien, C. 2002b. Host identification for three freshwater mussel species endemic to the
Altamaha River, Georgia. Ellipsaria 4(1):17.
O’Brien, C., and J. Brim-Box. 1999. Reproductive biology and juvenile recruitment of
the Shinyrayed Pocketbook, Lampsilis subangulata (Bivalvia: Unionidae) in the Gulf
Coastal Plain. American Midland Naturalist 142(1):129–140.
O’Brien, C., J. Williams, and M. Hoggarth. 2003. Morphological variation in glochidia
shells of six species of Elliptio from Gulf of Mexico and Atlantic Coast drainages in
the southeastern United States. Proceedings of the Biological Society of Washington
Ortmann, A.E. 1911. A monograph of the naiades of Pennsylvania, Parts I and II. Memoirs
of the Carnegie Museum 4(6):279–347, 4 plates.
Skelton, C.E., S. Cammack, and E. VanDeGenachte. 2002. Survey for Elliptio spinosa
(Altamaha Spinymussel) in the lower Ocmulgee River. Georgia National Heritage
Spooner, D.E., and C.C. Vaughn. 2006. Context-dependent effects of freshwater mussels
on stream benthic communities. Freshwater Biology 51:1016–1024.
Strayer, D.L., J.A. Downing, W.R. Haag, T.L. King, J.B. Layzer, T.J. Newton, and S.J.
Nichols. 2004. Changing perspectives on pearly mussels, North America’s most imperiled
animals. BioScience 54:429–439.
Strayer, D.L. 2008. Freshwater Mussel Ecology: A Multifactor Approach to Distribution
and Abundance. University of California Press, Los Angeles, CA.
Williams, J., M. Warren, Jr,, K. Cummings, J. Harris, and R. Neves. 1993. Conservation
status of freshwater mussels of the United States and Canada. F isheries 18(9):6–22.
Williams, J.D., A.E. Bogan, and J.T. Garner. 2008. Freshwater Mussels of Alabama and
the Mobile Basin in Georgia, Mississippi, and Tennessee. University of Alabama
Press, Tuscaloosa, AL.
Wisniewski, J.M., G. Krakow, B. Albanese. 2005. Current status of endemic mussels in
the lower Ocmulgee and Altamaha rivers. In K.J .Hatcher (Ed.). Proceedings of the
2005 Georgia Water Resources Conference. Institute of Ecology, Athens, GA. Available
online at http://www.gwri.gatech.edu/uploads/proceedings/2005/WisniewskiJGWRCpaper.
Zale, A., and R. Neves. 1982. Fish hosts of four species of lampsiline mussels (Mollusca:
Unionidae) in Big Moccasin Creek, Virginia. Canadian Journal of Zoology