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Preliminary characterization of digestive enzymes in freshwater mussels.

ABSTRACT Resource managers lack an effective chemical tool to control the invasive zebra mussel Dreissena polymorpha. Zebra mussels clog water intakes for hydroelectric companies, harm unionid mussel species, and are believed to be a reservoir of avian botulism. Little is known about the digestive physiology of zebra mussels and unionid mussels. The enzymatic profile of the digestive glands of zebra mussels and native threeridge (Amblema plicatd) and plain pocketbook mussels (Lampsilis cardium) are characterized using a commercial enzyme kit, api ZYM, and validated the kit with reagent-grade enzymes. A linear correlation was shown for only one of nineteen enzymes, tested between the api ZYM kit and a specific enzyme kit. Thus, the api ZYM kit should only be used to make general comparisons of enzyme presence and to observe trends in enzyme activities. Enzymatic trends were seen in the unionid mussel species, but not in zebra mussels sampled 32 days apart from the same location. Enzymatic classes, based on substrate, showed different trends, with proteolytic and phospholytic enzymes having the most change in relative enzyme activity.

KEY WORDS: freshwater mussel, zebra mussel Dreissena polymorpha, digestion, enzyme, Amhlema plicata, Lampsilis cardium


Since their introduction in the late 1980s, zebra mussels (Dreissena polymorpha Pallas, 1769) have spread across the United States and have negatively impacted native unionid populations (Strayer & Malcom 2007). While Dreissena polymorpha colonize and smother many unionid mussel species (Schloesser et al. 1996), they are also thought to potentially compete with unionid mussels for food resources (Baker & Levinton 2003). To address the competition issue, a method to comprehensively analyze the gut contents of zebra and unionid mussels is needed. Several studies have analyzed contents within the gut of mussels to identify what they are consuming (Yeager et al. 1994, Parker et al. 1998, Beck & Neves 2003, Mandal et al. 2007, Kamiyama 2011); gut content analysis does not differentiate between what is being digested and what will pass through a mussel. Results of gut content analyses have not provided conclusive evidence of dietary competition, leading some researchers to use stable isotope analysis to characterize potential dietary competition. When in fact, allowing more information about mussel diets than gut content analysis, factors including species, location, and time confound diet comparison. These confounding factors have led to a debate among researchers over mussel diet being composed of algae or bacteria. For example, Nichols and Garling (2000) reported that unionid mussels primarily feed on bacteria, whereas Baker and Levinton (2003) suggest they feed on algae. These two studies were conducted on mussel species that occupy different ecological niches, further adding to the uncertainty of mussel diets and potential for overlap. Thermally dependent metabolic rate (Hawkins 1985, Bosley et al. 2002, Gamboa-Delgado & Le Vay 2009) and the seasonal changes in food resources (Kreeger & Newell 2001, Taylor & Batzer 2010, Bergamino et al. 2011, Kamiyama 2011) both contribute to turnover of stable isotopes from the body tissue, further complicating stable isotope analysis. Stable isotopes thus provide valuable insight into the feeding habits of animals prior to collection, but not at the time they were collected.

Techniques to identify digestive enzymes are potentially more informative when characterizing mussel gut contents. Digestive enzymes are dependent on the types and chemical composition of the substrates present within the digestive tract (Deren et al. 1967, Rosensweig & Herman 1969, Nitsan et al. 1974. Kelly et al. 1991). Chemical composition of zooplankton and algae are highly variable (Raymont et al. 1971, Jagadeesan et al. 2010, Becker 2007). If multiple digestive enzymes could be analyzed at a single time, they would form a potential index to compare feeding and digestion among mussel species. To date, most studies using digestive enzymes in freshwater mussels have focused on the kinetics and absence or presence of one or more enzymes (Kuz'mina 1999, Areekijseree et al. 2002, Supannapong et al. 2008, Khrueanet et al. 2009, Palais et al. 2010, Golovanova 2011). Even though analysis of a very limited number of enzymes is important in the identification of optimal conditions and limitations to metabolism and the ability to digest specific substrates (e.g., protein, cellulose, etc.), it provides little information on the dietary content, particularly for comparisons among species. Analyzing a large suite of digestive enzymes concurrently may allow inference about dietary content and feeding status of an animal at the time of collection. A chromatic diagnostic enzyme kit (api ZYM) manufactured by bioMerieux Inc. allows for the simultaneous assessment of 19 digestive enzymes, which we proposed to use to compare the levels of digestive enzymes among freshwater mussel species.

Specific objectives include: (1) validate the use of api ZYM kit to measure and compare digestive enzyme profiles in freshwater mussels, and (2) determine if there are species-specific digestive enzymes. To the best of our knowledge, this is the first study to simultaneously measure multiple digestive enzymes from three species of freshwater mussels.


Any use of trade, product, or company name is for descriptive purposes only and does not imply endorsement by the U.S. Government.

Animals and Collection

Ten Amblema plicata, three Lampsilis cardium, and seven Dreissenapolymorpha were collected on August 9, 2010 in Pool 6 of the Mississippi River, near Winona, MN. On September 10, 2010, ten individuals of each species were collected from the same location. Water temperatures at the collection times were 28.6 and 19.4[degrees]C, respectively. After collection, digestive glands were excised from each A. plicata and L. cardium. Individual digestive glands were frozen on dry ice in the field and stored at -80[degrees]C in the laboratory until further processing. Whole D. polymorpha were also immediately frozen in the field and transported to the laboratory because magnification was required to extract their digestive glands. Upon thawing, D. polymorpha digestive glands were removed and processed identically to those from the A. plicata and L. cardium.

pH and Processing

Immediately after thawing, the pH of each digestive gland was determined with an AB15 pH meter and a Micro pH electrode (Accumet, Fisher Scientific, Fairhaven, NJ). Each digestive gland was then homogenized using a Microtube Pestle (USA Scientific, Inc., Ocala, FL). Samples were centrifuged at 23Xg for 20 min and supernatants retained for all subsequent analyses. De-ionized (DI) water was added to each sample to increase volumes and dilute the total protein. Total protein concentration was quantified using a microBCA assay (Thermo Scientific, Rockford, IL) according to the manufacturer's instructions on a Biotek Synergy 2 spectrophotometer (Biotek, Winooski, VT). Each sample was then diluted with DI water to 1.0 [micro]g protein/[micro]L.

Enzyme Analysis

Each sample was assayed for 19 digestive enzymes (Table 1) using api ZYM test kits (bioMerieux, Inc., Durham, NC) according to the manufacturer's instructions. The August samples required 2.5 [micro]g total protein loaded into each well of the test kit strip to provide adequate color production, whereas the September samples required 5.0 [micro]g total protein to produce a similar color change. Directly after the final step of the api ZYM assay, the contents of each well, including the blank, were transferred to a corresponding well in a 96-well plate. Absorbance of each well was measured at 400, 450, 500, 550, and 650 nm with a Synergy 2 MultiMode Microplate Reader (BioTek, Winooski, VT). Apparent enzyme activity (AEA) was determined at each wavelength using the following equation:

[AEA.sub.X] = (Abs[X.sub.sample]/[Ptn.sub.sample]) - (Abs[X.sub.blank]/[Ptn.sub.blank]),

where [AEA.sub.X] is the apparent enzyme activity determined by absorbance at wavelength X, AbsX is the absorbance of the sample/well at wavelength X, and Ptn is the amount of protein loaded into the well.

The appropriate wavelengths used to analyze each enzyme were determined using a standard curve and identifying the wavelength that produced a linear response curve. The standard curve was comprised of supernatant of the homogenate of a single Amblema plicata digestive gland in the following amounts of total protein: 0, 1.5, 3, 6, 9, or 12 [micro]g. Each strip was processed as mentioned above and absorbance measured at wavelength of 400, 450, 500, 550, and 600 nrn. Standard curves were developed for each wavelength and those wavelengths that provided the best fit model ([R.sup.2] [greater than or equal to] 0.90) were used for further analysis. These standards curves were used to calculate concentration from the absorbance of each enzyme assay. The data presented here are semiquantifiable, in that the data are accurate for relative comparisons of enzyme activity among samples, and should not be taken to indicate an absolute quantity. Those enzymes that did not provide an appropriate model were only considered present or absent.

Validation of Enzyme Kit (Absolute Quantification)

Using reagent-grade enzymes, we evaluated if enzyme activity determined with the api ZYM kit could be quantified. Alkaline phosphatase, acid phosphatase, leucine arylamidase, and jV-acteyl-P-glucosaminidase (NAGase) were the only enzymes available for analysis. Each enzyme was serially diluted: alkaline phosphatase: 1.0, 2.0, and 3.0 mU/well; acid phosphatase: 0.25, 0.50, and 0.75 mU/well; leucine arylamidase: 0.75, 1.0, and 2.0 mU/well; NAGase: 3.0, 4.0, and 5.0 mU/well. All enzymes were purchased from Sigma-Aldrich (St. Louis, MO). Absorbances at the targeted wavelengths were measured for the contents of the well that corresponded to the targeted enzymes.


A t-test was used to determine if shell length of mussels, within a species, differed between sampling times. Enzyme concentrations were compared among species by analysis of variance (ANOVA) and Tukey's post hoc test from samples collected in September. August samples were not analyzed owing to low number of Lampsilis cardium samples. T-tests were used to identify changes in enzyme concentrations between August and September using data from Amblema plicata and Dreissena polymorpha. Lampsilis cardium were excluded from this analysis owing to limited sample sizes. Enzymes that were not quantified were simply expressed as present or absent. Statistical analyses of differences in digestive enzymes among species were performed using SYSTAT 11.0 (Systat Software, Inc., San Jose, CA) with a significance level of P < 0.02. This significance level was used owing to limited sample sizes and to minimize Type II errors. Statistical analyses for comparing the absolute quantification data were performed using R version 3.1.0 with a significance level of P < 0.05. Statistical analysis for shell length data between sampling dates was performed using R version 3.1.0 with a significance value of P < 0.02.


Body Size and pH

Within each species, no differences in shell lengths were observed between sample times. The mean (SD) shell length for Amblema plicata was 82.0 (14.0) mm in August and 75.1 (5.7) mm in September (P = 0.176). Mean lengths of Lampsilis cardium were 123.8 (6.5) mm and 108.6 (5.9) mm in August and September, respectively (P = 0.034). In August, Dreissena polymorpha were 20.4 (2.0) mm and 20.8 (2.6) mm long in September (P = 0.706).

Mean pH (SE) of digestive glands were generally circum-neutral and did not differ between months for each species (P values: Amblema plicata: 0.994; Lampsilis cardium: 0.970; Dreissena polymorpha'. 0.980). The pH of the digestive glands were 6.753 (0.046) in A. plicata, 6.881 (0.048) in D. polymorpha, and 7.003 (0.067) in L. cardium. Significant differences were observed in the pH of the digestive glands of the three mussel species. The pH of digestive glands from D. polymorpha did not differ from those in A. plicata (P = 0.234) or L. cardium (P = 0.254), but digestive gland pH was significantly different in A. plicata compared with L. cardium (P = 0.007).

Validation of Enzyme Kit

Only seven enzymes were found to produce a positive linear trend in one of the wavelengths measured (Table 1). Six enzymes were semiquantifiable at 550 nm and naphthol-AS-BI-phosphohydrolase was quantifiable at 600 nm. Enzymes that were semiquantifiable were phospholytic (alkaline phosphatase, acid phosphatase, and naphthol-AS-BI-phosphohydrolase), glyocolytic ([beta]-galactosidase, NAGase, fucosidase) or proteolytic (leucine arylamidase). No lipolytic enzymes (lipase, esterase, and esterase lipase) could be quantified. Twelve enzymes were classified as present or absent.

Of the seven enzymes found to have a positive linear response with the Amblema plicata digestive gland homogenate, reagent-grade enzymes were used to generate a standard curve of the absorbance versus enzyme concentration. A comparison of the resulting api ZYM-generated concentrations and the concentrations generated via the specific enzyme kits showed that of the four enzymes tested, only one resulted in a significant linear correlation, although there was still substantial scatter among data points (data unpublished; NAGase, P < 0.05; Fig. 1).

Comparison Among Species

In general, concentrations of the seven quantifiable digestive enzymes differed significantly between native mussels and zebra mussels, but most enzymes were similar between the two native mussel species. Of the enzymes that had differences, most were phospholytic enzymes and will be the primary focus because of the lack of knowledge on the other quantifiable enzymes. Values presented here are based upon standard curve analysis with reagent-grade enzymes; values in figures are merely for graphical representation. There was a greater quantity of alkaline phosphatase in Amblemci plicata than in Lampsilis cardium, 0.826 (0.129) units/[micro]g total protein and 0.334 (0.075) units/pg total protein (P = 0.02), respectively (Fig. 2). No differences were determined in alkaline phosphatase (Fig. 2) quantity between Dreissena polymorphci and A. plicata or L. cardium. In D. polymorpha, leucine arylamidase (Fig. 3; 0.623 (0.046) units/pg total protein) was greater than that in either A. plicata or L. cardium (P [less than or equal to] 0.01). Also, acid phosphatase (Fig. 2) concentrations in D. polymorpha [1.525 (0.125) units/[micro]g total protein] were greater than that determined in A. plicata [0.960 (0.091) units/[micro]g total protein] or L. cardium [0.769 (0.050) units/pg total protein; P < 0.01]. All other semiquantifiable enzymes (Figs. 2-4) were similar among species. The enzymes classified as absent or present were identified in all three species.

Comparison Between Sample Times

In general, concentrations of digestive enzymes differed between the August and September sampling dates. Concentrations of phospholytic enzymes were significantly less in Amhlema plicata captured in September than those captured in August, but no differences were determined for those enzymes in Dreissena polymorpha (Fig. 2). Concentrations of naphthol-AS-BI-phosphohydrolase in A. plicata decreased from 12.990 (2.170) units/pg total protein in August to 0.934 (0.621) units/pg total protein in September (P<0.01). Acid phosphatase decreased from 3.820 (0.659) to 0.960 (0.289) units/[micro]g total protein and alkaline phosphatase decreased from 2.937 (1.254) to 0.825 (0.407) units/[micro]g total protein from August to September, respectively (P < 0.01). The glycolytic enzyme NAGase (Fig. 4) did not change in concentration between August and September in either A. plicata or D. polymorpha {P = 0.04). Both fucosidase and [beta]-galactosidase (Fig. 4) decreased in concentrations from August to September in A. plicata and D. polymorpha (P < 0.01). The concentration of leucine arylamidase (Fig. 3) was similar in A. plicata captured in August and those captured in September (P = 0.07). Leucine arylamidase increased from 0.206 (0.097) to 0.623 (0.144) units/[micro]g total protein from D. polymorpha captured in August and September (P < 0.01).


The study of digestive enzymes provides insight into the diet and physiology of an animal, which may guide programs developing tools for the management of invasive species. The api ZYM kit was useful for the determination of the absence/ presence of 19 digestive enzymes in three species of mussels. However, of these enzymes, only seven were significantly correlated with the amount of digestive enzyme and thus could be quantified. These seven assays can be useful for making accurate comparisons of enzyme activity. The remaining 12 enzymes can be used for absence/presence detection, and thus identify those enzymes present, which require additional analysis.

The following discussion on increased enzyme activity in Dreissena polymorpha should be interpreted with caution. As a result of Dreissena polymorpha being dissected differently than unionid mussels, the concentration of enzymes used in the analysis could be lower in D. polymorpha owing to dilution by other cellular components and tissues. The majority of enzymes analyzed by the api ZYM enzyme test kit are primarily located in tissues associated with digestive function (BRENDA enzyme portal; accessed September 7, 2013; Thus, the approach used for sampling D. polymorpha has likely underestimated their digestive enzyme activity.

The potential for absolute quantification was investigated to determine whether a linear response could be found between the concentrations generated for each enzyme between the two assay kits (unpublished data). The data indicated that only NAGase had a linear relationship, albeit weak, when analyzed with both the digestive gland homogenate and the reagent-grade enzyme. Other enzymes had linear responses with the reagent-grade enzymes. However, the assay was saturated at very low amounts of the enzyme (data not shown). Our results suggest that the api ZYM kit should not be used to quantify enzymatic activity, but could be a valuable tool to compare relative activities among samples and identify trends.

Even though all of the enzymes analyzed were present in all species, the activities of the enzymes in unionids appeared to be greater than those in Dreissena polymorpha in August, indicating unionid mussels may be using a different food source than D. polymorpha at that time. This conclusion contrasts with other studies that used gut contents and stable isotopes to analyze mussel diets and found they feed upon similar food sources. However, these techniques are limited in determining the actual food source, and stable isotope methods are unable to easily discriminate the signals of bacteria, picoalgae, and fungi.

The size of the food particles and the method of filtration also influence the mussels' choice of food source. Unionid mussels and Dreissena polymorpha have been shown to have an overlapping range in the size of food particles they can filter from the water column (Vanderploeg et al. 1995, Baker & Levinton 2003, Dionisio Pires et al. 2004). While Dreissena polymorpha have been shown to have a larger gill surface area (per mg dry tissue) than unionid mussels, it has also been shown that the number of cilia per cirri is similar between unionid mussels and D. polymorpha (Silverman et al. 1995, Silverman et al. 1997). Although filtration processes do not directly impact enzymatic activity, they do impact the types of food resources available for digestive activity. Determining the types of food resources retained by filtration processes and determining the digestive enzyme activity may provide greater clarity into the mussel diet at the time of collection.

Digestive enzyme activity, like other physiological, biochemical, and metabolic rates, are often temperature dependent. Spooner and Vaughn (2008) found differences in rates of resource acquisition and assimilation in freshwater mussels among different thermal regimes. In the current study, the significant differences in digestive enzyme activity between August and September may have resulted from the 10[degrees]C cooler temperatures in September. Ribbed mussels Geukensia demissa have been reported to have diet shifts between summer/fall and winter/spring (Kreeger & Newell 2001). Other species of ectothermal animals, such as Farfantepenaeus duorarum and Sparus aurata, have been shown to have seasonal differences in digestive enzyme activity (Aragon-Axomulco et al. 2012, Sanchez-Muros et al. 2013). Because metabolic functions are temperature dependent and vary among species, seasonal differences in enzymatic activity may also exist among freshwater mussels (Fanslow et al. 2001, Spooner & Vaughn 2008, Pandolfo et al. 2010, Ganser et al. 2013). Seasonal trends of metabolic activity have been described in marine molluscan literature. For example, in Crassostrea virginica phosphofructokinase and pyruvate kinase change from catabolic function in winter to anabolic function in summer (Greenway & Storey 2000). Changes in kinetic properties of phosphofructokinase and pyruvate kinase in the tissues of Littorina littorea have also been observed (Greenway & Storey 2001). In Modiolus modiolus, the concentration of rate-limiting metabolic enzymes increased to compensate for the loss of activity due to low temperatures, and in Mytilus galloprovincialis there was significant difference in pyruvate kinase activity from the mantle tissue between winter and spring months (Lesser & Kruse 2004, Ioannou et al. 2009).

Of the 19 enzymes we assayed, only leucine arylamidase levels did not differ between months in unionid mussels, but did change in Dreissena polymorpha. Leucine arylamidase has been linked with nitrogen excretion in the blue mussel Mytilus edulis (Hilbish & Koehn 1985). Blue mussels with the [Lap.sup.94] allele excreted more nitrogen than those lacking the [Lap.sup.94] allele (Hilbish & Koehn 1985). Typically, Dreissena polymorpha are most metabolically active, as measured by oxygen consumption/nitrogen excretion, in fall (Sprung 1995). In contrast, unionid mussels are typically most metabolically active, as measured by oxygen consumption/nitrogen excretion, in spring and summer (Baker & Hornbach 2001). This difference in metabolic activity may account for the increase in enzyme activity in D. polymorpha and the decrease in enzyme activity in unionid mussels in this study. Although comparisons between D. polymorpha and unionids should be viewed cautiously for the reasons listed above, we are not aware of any data on enzyme levels in other freshwater molluscs for comparison.

In conclusion, the api ZYM enzyme test kit was determined to be a valuable tool in determining trends in enzyme activity and comparing relative digestive enzyme activities among species. Our data suggests, however, that specific enzyme kits should be used to determine absolute enzyme quantification. Our results also suggest that Dreissena polymorpha and unionid mussels may feed on different food sources even if they occupy similar trophic positions. These data further suggest that native mussel feeding rates may decrease in fall even if D. polymorpha continue to feed. Additional data on seasonal patterns in digestive enzymes in freshwater molluscs may help develop additional control strategies for D. polymorpha. For example, a recently approved product for controling D. polymorpha within defined discharge and open water systems (Zequanox) causes necrosis of the digestive gland and stomach epithelial cells (Molloy et al. 2013). Additional work to characterize the digestive enzymes present in the gastrointestinal tracts of freshwater molluscs, seasonal variation in enzyme presence and activity, and the effect of those enzymes on Zequanox or other potential controls may help create formulations which maximize activity in dreissenid mussels whereas minimizing potential impacts on native species. Research on the use of digestive enzymes to inform the development of control tools is in progress, such as the identification of differences in leucine arylamidase activity between D. polymorpha and native mussels in our study. Our research was able to find that D. polymorpha leucine arylamidase may present one enzyme managers can exploit to develop a control tool.


This work was supported by the U.S. Environmental Protection Agency Great Lakes Restoration Initiative and the USGS Science Support Program. We thank Jim Luoma, Alissa Ganser, Patricia Ries, and Ashley Hunt for their assistance with mussel collections, and we thank Shelby Storsveen for her help with enzymatic analysis.


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(1) Biology Department, University of Wisconsin--La Crosse, 1725 State Street, LaCrosse, WI 54601; (2) U.S. Geological Survey, Upper Midwest Environmental Sciences Center, 2630 Fanta Reed Road, La Crosse, WI 54603; (3) Chemistry and Biochemistry Department, University of Wisconsin-La Crosse, 1725 State Street, La Crosse, WI 54601

* Corresponding author. E-mail:

DOI: 10.2983/035.034.0225

Enzymes assayed in the api ZYM kit and the wavelength at
which the absorbance was measured (nm) if a linear
response was observed.

                         wavelength      Trend line
Enzyme                      (nm)           equation       [R.sup.2]

Blank                        --               --
Alkaline phosphatase *       550       y = 0.04533.x -      0.973
Esterase                     --               --             --
Esterase lipase              --               --             --
Lipase                       --               --             --
Leucine arylamidase *        550        y = 0.0441x -       0.950
Valine arylamidase           --               --             --
Cystine arylamidase          --               --             --
Trypsin                      --               --             --
[alpha]-chymotrypsin         --               --             --
Acid phosphatase *           550        y = 0.0432x -       0.964
Naphthol-AS-BI-              600        y = 0.0137x +       0.910
  phosphohydrolase *                        0.0122
[alpha]-Galactosidase        --               --             --
[beta]-Galactosidase *       550        y = 0.0774x -       0.959
[beta]-Glucuronidase         --               --             --
[alpha]-Glucosidase          --               --             --
[beta]-Glucosidase           --               --             --
N-acetyl-p-                  550        y = 0.1008x -       0.980
 glucosaminidase *                         0.0087
Mannosidase                  --               --             --
Fucosidase *                 550        y = 0.0907x -       0.993

The validation of the quantification of the api ZYM kit at these
wavelengths resulted in the following trend line equations.

* Enzymes analyzed beyond a presence/absence basis.
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Author:Sauey, Blake W.; Amberg, Jon J.; Cooper, Scott T.; Grunwald, Sandra K.; Newton, Teresa J.; Haro, Rog
Publication:Journal of Shellfish Research
Date:Aug 1, 2015
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