Variation in Glycogen Distribution among Freshwater Bivalve Tissues:
Simpliﬁed Protocol and Implications
a and Karel Douda*
Department of Zoology and Fisheries, Czech University of Life Sciences Prague, Kam
a 129, CZ-16500, Prague,
Glycogen is a primary metabolic reserve in bivalves and can be
suitable for the evaluation of bivalve condition and health status,
but the use of glycogen as a diagnostic tool in aquaculture and
biomonitoring is still relatively rare. A tissue biopsy combined with
a simpliﬁed phenol–sulfuric acid method was used in this study to
evaluate the inter- and intraindividual variation in the glycogen con-
centrations among several tissues (foot, mantle, gills, adductor mus-
cle) of the unionid bivalve, the duck mussel Anodonta anatina. This
short report documents that individual bivalves differ in the spatial
distribution of glycogen among tissues. Sampling of different types
of tissues can cause distinct results in the evaluation of energetic
reserves at the individual level. At the same time, spatial variability
in glycogen content has the potential to provide a more detailed
evaluation of physiological conditions based on tissue-speciﬁc glyco-
gen storage. The results obtained and the simpliﬁed methodology
provide a new opportunity for researching the energetic reserves and
health status of freshwater mussels in various applications.
Monitoring the energetic reserves of aquatic inverte-
brates is still rarely implemented, although invertebrates
are increasingly being used in the food production sector,
in conservation aquaculture, and for biomonitoring (Koop
et al. 2008). The evaluation of the condition of macroin-
vertebrates is often determined indirectly from growth and
mortality data, but more speciﬁc physiological markers
can be critical for the early identiﬁcation of changes in
their health status (Fritts et al. 2015b).
This study addresses freshwater mussels, order Union-
ida, which are increasingly being propagated in aquaculture
facilities for conservation purposes, and there is a need to
employ reliable and noninvasive methods to assess their
energetic status (Fritts et al. 2015a). Despite the critical
conservation status of freshwater mussels, the determina-
tion of the physiological condition of this group is mostly
based on whole-body analyses performed by lethal methods
(Gustafson et al. 2005). These methods include the analysis
of the lipid and fatty acid composition in adult mussels
(Prato et al. 2010) and the analysis of lipids or glucose in
juveniles (Tankersley 2000; Sim-Smith and Jeffs 2011). In
contrast, nonlethal methods can be applied for a larger part
of a population to obtain more replications (Gustafson
et al. 2005) and for the evaluation of changes during long-
term studies. Nonlethal methods are particularly important
for the research of threatened and endangered species
(Naimo et al. 1998; Fritts et al. 2015a), where the sacriﬁce
of individuals could put populations of such species in jeop-
ardy (Naimo et al. 1998; Haskell and Pan 2010).
Glycogen is the primary metabolic reserve in mussels
(Ke and Li 2013). Stored glycogen is a source of glucose,
which can be mobilized to tissues (Martinez-Pita et al.
2012). Glycogen level is suitable as a physiological marker
for the evaluation of the condition and health of bivalves
(McGoldrick et al. 2009; Sim-Smith and Jeffs 2011; Cor-
deiro et al. 2017). Glycogen quickly reacts to changes in
the environment (Fisher and Dimock 2006), and it is
connected to the nutritional condition, different types of
stress, stages of the life cycle, and sexual maturity
(de Zwaan and Zandee 1972; Dridi et al. 2007; Anacleto
et al. 2013; Cordeiro et al. 2017). Monitoring changes in
glycogen levels has been used in various studies dealing
with bivalves, for example, to monitor stress (Fearman
and Moltschaniwskyj 2010; Fritts et al. 2015b; Andrade
et al. 2017) under different conditions such as starvation
(Cordeiro et al. 2016) or transportation (Yusufzai et al.
*Corresponding author: firstname.lastname@example.org
Received June 8, 2018; accepted December 5, 2018
Journal of Aquatic Animal Health 31:107–111, 2019
©2019 American Fisheries Society
ISSN: 0899-7659 print / 1548-8667 online
2010; Anacleto et al. 2013; Cordeiro et al. 2017), or in
ecotoxicological studies (Hazelton et al. 2014).
The biopsy of foot tissue followed by the phenol–sulfu-
ric acid method of glycogen determination by Naimo
et al. (1998) is a nonlethal method for the analysis of this
physiological marker in freshwater mussels. Glycogen can
be determined by means of the glycogen biopsy method
developed by Naimo et al. (1998) and is a very promising
marker of the physiological condition in bivalves. How-
ever, it is not routinely applied in aquaculture and conser-
vation physiology, probably because of the high workload
and material consumption of this method and of the lim-
ited knowledge of the spatial and temporal dynamics of
glycogen reserves in freshwater bivalve tissues.
The aim of this study was to determine the distribution
of glycogen in soft-body tissues (foot, gills, mantle, and
adductor muscle) in a freshwater bivalve, the duck mussel
Anodonta anatina, to estimate the potential effect of glyco-
gen spatial distribution on the evaluation of the energetic
status of a bivalve. We also aimed to simplify the glycogen
analysis method (phenol–sulfuric acid method) by Naimo
et al. (1998) to reduce the workload, costs, and material
consumption for easier ampliﬁcation of the methodology.
The duck mussel was used as a test organism because it
is a widespread European species (Lopes-Lima et al. 2017)
and has a potential use in biomonitoring (Falfushynska
et al. 2013). Six adult individuals (shell length, 82–
98.5 mm) sampled in the Lu
znice River, Czech Republic
(49°18024″N, 14°30014″E), were transported in a 25-L tank
with aerated (through an airstone) river water to the labo-
ratory where the tissue sampling was performed on the
same day (November 6, 2017). Biopsy samples were taken
through gently opened valves (using a shell opening
device) from the outer edge of the foot (posterior part),
mantle (medial part), and gills (medial part). Dissecting
scissors were used to cut off three independent samples
close to each other from the foot, mantle, and gill tissue,
in that order, from each individual. Then, the individual
was sacriﬁced by cutting the adductor muscles, and the
shell was opened completely. The tissue of the anterior
adductor muscle was sampled in the same way as that for
other tissues. The target weight of samples was 4–10 mg
(sample weight identiﬁed as suitable for nonlethal biopsy
of foot tissue in Unionidae and glycogen analysis protocol
by Naimo et al. 1998), which corresponded to a piece of
tissue approximately 6 9191.5 mm in size. There were
68 samples collected (four pairs of replicated samples were
merged due to the small sample weight), and the true
weight of the ﬁnal samples ranged from 2.17 to 12.68 mg
(mean weight, 7.36 mg) of wet tissue. Samples were stored
at 75°C and processed within 40 d.
The method by Naimo et al. (1998) was used with sev-
eral modiﬁcations (details may be found online in the Sup-
porting Information section at the end of the article), and
the bias associated with glycogen determination was esti-
mated by recovery of known additions using matrix stan-
dards. The precision of the method was estimated from
triplicate analyses of all matrix standards (relative stan-
dard deviation [RSD]); the method detection limit (MDL)
was determined using 11 blank samples from three differ-
ent analysis sets according to the EPA (2016) guidelines.
Calibration standards.—Glycogen calibration standards
were prepared by dissolving 40 mg of powdered oyster
glycogen (type II, Sigma-Aldrich, St. Louis, Missouri) in
20 mL of deionized water and then creating serial dilu-
tions of 2,000, 1,000, 500, 250, and 125 mg/L immediately
Internal standards.—The in-house reference material
was prepared according Naimo et al. (1998) from the
homogenized foot tissue of six duck mussels sampled on
July 24, 2017, from the same location as the study samples.
Two milliliters of 30% KOH (Penta, Prague, Czech Repub-
lic) was added per gram of tissue to a 3-mL cryovial (Sim-
port, Beloeil, Quebec), heated for 20 min in a water bath at
100°C (RTC Basic, IKA, Wilmington, North Carolina),
then vortexed for 10 s (MS2 Minishaker, IKA), and stored
in a freezer at 75°C. The in-house reference material was
thawed at room temperature and vortexed before use during
the whole study (four analytical days).
Spiked calibration standards.—Spiked samples were pre-
pared the same way as the aqueous calibration standards,
but they were spiked by adding 10 μL of the in-house refer-
Digestion and extraction of glycogen.—Glycogen from
all standards and samples was digested and extracted from
tissues. Thirty percent KOH was added to the samples in
3-mL cryovials in a volume of 100 μL to the tissue sam-
ples, blank solution, and in-house reference material sam-
ples. The volume of 30% KOH added to the spiked
samples and aqueous calibration standards was 280 μL.
Then, cryovials were boiled in a water bath for 20 min for
homogenization. In the next step, 96% ethyl alcohol
(Penta) was added to the solution, and cryovials were
placed in a boiling water bath for 15 min. The volume of
ethyl alcohol was 1.5 times more than the added KOH to
prevent the precipitation of other polysaccharides (Naimo
et al. 1998). The solutions were diluted with deionized
water to the same total solution volume; after homoge-
nization, 2,660 μL of solution were removed by pipette
from the samples, and 390 μL of deionized water were
added to the solutions to reach the optimal ratio of water
A AND DOUDA
volume in the analyzed samples (see the detailed step-by-
step protocol of the modiﬁed methodology 1–13 in the
Quantiﬁcation of glycogen.—Quantiﬁcation of glycogen
was based on spectrophotometry. Forty microliters of 80%
phenol (Carl ROTH, Karlsruhe, Baden-W€
Germany) and 2,180 μL of 96% sulfuric acid (Penta) were
added to the sample solutions to gain coloration. A 250-μL
aliquot of the solution of each sample was pipetted into a
96-well microplate (Anicrin S.R.L., Scorz
e, Venice, Italy),
and the absorbance of the samples was determined by
spectrophotometry (see the detailed step-by-step protocol
of the modiﬁed methodology 14–19 in the Supplement
available in the online version of this article).
The content of glycogen in samples was determined by
using the calibration slope, which was calculated from the
absorbance of triplicate samples of the aqueous calibration
standard of used concentrations. The calibration slope was
estimated individually for every analytical set with linear
regression models. The mean recovery of glycogen in
spiked samples, the mean CV (100SD/mean) (SD) for
each concentration of aqueous calibration standards, and
the mean percentage difference between slopes were calcu-
lated according to Naimo et al. (1998) from three repli-
cated samples on four analysis dates.
A two-way ANOVA was used to examine the effects of
tissue (foot, mantle, gills, adductor muscle) and the mussel
individual (A–F) on the glycogen content. The interaction
term of explanatory variables was included in the analysis.
A post hoc Tukey’s honestly signiﬁcantly different (HSD)
test was used to identify the signiﬁcant differences between
pairs of tissues. Before statistical analyses, data were
assessed for the homogeneity of variance and normality
using Levene’s test and a Kolmogorov–Smirnov test,
respectively. Glycogen data were transformed by means of
a natural-log transformation before analyses. The data
analysis was performed using the R software (R Develop-
ment Core Team 2017).
The glycogen concentrations in duck mussels ranged
from 5.6 mg to 60.1 mg/g wet tissue, with a mean concen-
tration of 20.7 mg/g (SD, 10.7). The mean glycogen con-
tent in the biopsied samples was 0.15 mg (SD, 0.07), and
MDL was established to be 0.049 mg of glycogen in the
whole analyzed sample (0.0075 mg of glycogen in the ﬁnal
solution for spectrophotometry).
The slopes of the calibration standard and spiked cali-
bration standard regression lines were not signiﬁcantly dif-
ferent (ANOVA: P>0.05, with the interaction term of
concentration level by spiking) in all analytical sets,
validating the use of the calibration standards line to pre-
dict the glycogen content. The aqueous calibration standard
curves had a mean slope of 258.28 (range, 213.44–283.44),
mean intercept of 0.1973 (range, 0.1860–0.2113), and mean
of 0.9769 (range, 0.9403–0.9928). The curves of the
spiked samples had a mean slope of 261.55 (range, 209.35–
290.57), mean intercept of 0.2889 (range, 0.2601–0.3058),
and mean R
of 0.9490 (range, 0.9275–0.9669). The mean
percentage difference between slopes was 1.258%.
The mean recovery of glycogen in spiked samples was
85% (SD, 13) for the concentration of 125 mg/L of the
aqueous calibration standard, 82% (SD, 27) for a concen-
tration of 500 mg/L, 109% (SD, 20) for a concentration of
1,000 mg/L and 83% (SD, 25) for a concentration of
2,000 mg/L. The CVs of these concentration samples were
7.7% (SD, 2.7), 10.5% (SD, 3.4), 7.4% (SD, 4.5) and 7.9%
(SD, 1.6), respectively.
The glycogen concentration signiﬁcantly differed among
tissues (two-way ANOVA: F
=6.9, P<0.001; Figure 1).
The mean glycogen concentration was 18.7 mg/g (SD, 4.9)
in the foot tissue, 20.1 mg/g (SD, 16.1) in the mantle,
20.2 mg/g (SD, 8.6) in the adductor mussel, and 26.1 mg/g
(SD, 9.0) in the gills (Figure 1; Table S1 available in the
Supplement). Subsequent post hoc comparison tests revealed
higher glycogen content in gills than in all other tissues (all
There was a signiﬁcant effect of individual mussels
(two-way ANOVA: F
=22.0, P<0.001; Figure 1). A
signiﬁcant interaction term between tissue and specimen
(two-way ANOVA: F
=3.3, P<0.01) demonstrated
that individual mussels differed in the spatial distribution
of glycogen among tissues.
Foot Gills Mantle Adductor
FIGURE 1. Glycogen content in duck mussel tissues sampled in the
znice River, Czech Republic. The mean (black line), median (white
line), interquartile range, minimum–maximum (without values >1.59
interquartile range) are displayed by box plots. Values measured for
individual mussels (six individuals, two or three samples per tissue) are
indicated by different symbols.
A simpliﬁed methodology for glycogen analysis was
demonstrated to be precise and reliable for the identiﬁca-
tion of differences in the glycogen level between types of
tissue in the duck mussel as well as in the body distribution
of glycogen between individuals. The results conﬁrm previ-
ous ﬁndings (de Zwaan and Zandee 1972; Naimo and
Monroe 1999) that there is a signiﬁcant difference in glyco-
gen levels in different types of tissues of unionid bivalves
and that the glycogen content in the mantle is the least
stable, while the most stable glycogen is in foot tissue (Fig-
ure 1). Furthermore, this study demonstrated that (despite
relatively high variability even within the same anatomical
structure) individual mussels differed signiﬁcantly in the
body distribution of the glycogen content. A tissue-speciﬁc
glycogen evaluation can provide more detailed data for the
monitoring the health and condition of mussels and can
provide new valuable information for future sampling,
where more than one type of tissue for the glycogen analy-
sis can be quantiﬁed. These ﬁndings also provide a new
view of the evaluation of results from ﬁeld studies where
only one tissue was sampled. Although the duck mussels
were not sampled nonlethally in this study, previous
ﬁndings corroborate that both foot and mantle tissue can
be biopsied nondestructively and without increasing
mortality in a long-term perspective (Berg et al. 1995;
Naimo et al. 1998). However, the effect of gill or adductor
mussel biopsy (or simultaneous extraction of more samples
per individual) on the survival rate of unionid mussels
needs to be tested.
The modiﬁed method is more economical by reducing
the analytical supplies, speciﬁcally the sulfuric acid volume
by 56.4%, and by using only one 3-mL cryovial per sam-
ple instead of transferring the solution among three types
of laboratory test tubes. The analysis is easily performed
by one person without assistance, and it allows an analysis
of up to 60 samples (tissue samples plus calibration of
standard solutions) in one analytical set. These improve-
ments will signiﬁcantly reduce amount of hazardous waste
that is produced by this analysis.
Further studies are needed to clarify the connection
between the glycogen level and its distribution in the body,
considering changes that occur annually (de Zwaan and Zan-
dee 1972), as well as other factors. Bivalves are exposed to
annual cycles of food availability which highly inﬂuences
their glycogen reserves (Albentosa et al. 2007; Cordeiro et al.
2016). Glycogen storage and utilization are also closely con-
nected to the annual reproductive cycle (Lemaire et al. 2006)
because glycogen reserves can be converted into lipids during
the mating season for gamete development (Martinez-Pita
et al. 2012; Ke and Li 2013; Irisarri et al. 2015). Glycogen is
catabolized to add glucose to hemolymph when the glucose
level declines or when it is mobilized by the inﬂuence of some
stress factor (Fritts et al. 2015a). The glycogen concentrations
in bivalve soft tissues are largely used for monitoring the
impacts of stress under different conditions (Anacleto et al.
2013; Cordeiro et al. 2016, 2017) and in ecotoxicologal stud-
ies (Hazelton et al. 2014), because the glycogen level
decreases long before changes in growth and survival rates
are known (Sim-Smith and Jeffs 2011). The simpliﬁcation of
the methodology used in this study allows this method to be
used for routine applications and highlights the importance
of tissue-speciﬁc analyses for understanding mussel energetic
metabolism. In particular, it would be promising to focus on
the link between speciﬁc types of tissues and the condition of
the individual and its environment during different periods of
the year and with respect to environmental conditions. This
can help determine the reasons for the variation in the distri-
bution of glycogen in the body so that glycogen evaluations
can be put to practical use. Regarding the pilot character of
this study, further data are needed to establish which organ
and at which life stage and gender of the mussels is the best
for nonlethal biomonitoring.
In summary, using the simpliﬁed nonlethal glycogen
determination method, this study documented that indi-
vidual freshwater duck mussels differ in their spatial distri-
bution of glycogen among tissues. The results obtained
and the simpliﬁed methodology provide new opportunities
for the research of energetic reserves and the health status
of freshwater mussels in various aquaculture and conser-
This study was supported by the Czech Science Foun-
dation (19-05510S) and European Regional Development
Fund (Project No. CZ.02.1.01/0.0/0.0/16_019/0000845).
There is no conﬂict of interest declared in this article.
Karel Douda https://orcid.org/0000-0002-7778-5147
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Additional supplemental material may be found online
in the Supporting Information section at the end of the