Trophic Position and Potential Food Sources of 2 ...

Trophic Position and Potential Food Sources of 2 Species of Unionid Bivalves (Mollusca:Unionidae) in 2 Small Ohio Streams Author(s): Alan D. Christian, Brittany N. Smith, David J. Berg, James C. Smoot and Robert H. Findlay Source: Journal of the North American Benthological Society, Vol. 23, No. 1 (Mar., 2004), pp. 101-113 Published by: Society for Freshwater Science Stable URL: http://www.jstor.org/stable/1468400 . Accessed: 27/08/2013 14:19 Your use of the JSTOR archive indicates your acceptance of the Terms & Conditions of Use, available at . http://www.jstor.org/page/info/about/policies/terms.jsp

. JSTOR is a not-for-profit service that helps scholars, researchers, and students discover, use, and build upon a wide range of content in a trusted digital archive. We use information technology and tools to increase productivity and facilitate new forms of scholarship. For more information about JSTOR, please contact [email protected].

.

Society for Freshwater Science is collaborating with JSTOR to digitize, preserve and extend access to Journal of the North American Benthological Society.

http://www.jstor.org

This content downloaded from 130.160.104.221 on Tue, 27 Aug 2013 14:19:28 PM All use subject to JSTOR Terms and Conditions

J. N. Am. Benthol.Soc.,2004,23(1):101-113 ? 2004by The North AmericanBenthologicalSociety

Trophic position and potential food sources of 2 species of unionid bivalves (Mollusca:Unionidae) in 2 small Ohio streams ALAN D. CHRISTIAN1 AND BRITTANY N. SMITH2 Departmentof Zoology, PearsonHall, Miami University, Oxford, Ohio 45056 USA DAVID J. BERG3 Departmentof Zoology, Miami University, 1601 Peck Boulevard,Hamilton, Ohio 45011 USA JAMES C. SMOOT4 AND ROBERT H. FINDLAY5 Departmentof Microbiology,PearsonHall, Miami University, Oxford, Ohio 45056 USA Abstract. We determinedthe trophicpositions of 2 species of freshwatermussels, Elliptiodilatata and Ptychobranchus fasciolaris,from 2 small streams in central Ohio by measuring stable C and N isotope ratios and digestive fluid enzyme activities. We also examined stable C and N isotopes, microbialbiomass, microbialcommunity structure,nutrient (i.e., C, N, and P) concentrations,and contributionof microbialC to total fine particulateorganic C (FPOC).We hypothesized that 1) allochthonousinputs compose most of FPOC,2) mussels use fine particulateorganicmaterial(FPOM) as a food source, and 3) mussels respond to the low-protein content of FPOMby showing high proteaseactivity.MicrobialC composed35 to 86%of total FPOCduringthe autumnsamplingperiod. FPOMstable isotope values varied seasonally,whereas 813C and 815N content in mussel tissue was spatially (i.e., among sites) and temporallysimilar.Musselswere 2 to 4%omore depletedin 813Cthan seasonal FPOM.Digestive fluid enzymes were spatially and temporallystable across species, with activity of esterase > protease > lipase > glucosidase.Lipase:proteaseof digestive fluids from mussels were 1, whereas detritivorous species had ratios 50%). Fish and mussel diversity in the Big Darby Creek system is high (100 and 41 species, respectively; see Trautman 1981 and Watters 1990) with high water quality (Ohio EPA 1987).

Sestonand musselsampling Three 4 L water column samples were collected in summer (July 1998) at BD1 (for seston) and in autumn (October 1998) at BD1, BD2 and LD1 (seston, microbial biomass, and community structure). Water samples were filtered through a 250 pLmmesh to remove large, inedible parti-


TROPHICPOSITIONOF MUSSELS

2004]

cles (Wissing 1997) before being stored in acidwashed Nalgene jars. To collect FPOM, 100 to 500 mL of filtrate were passed through preweighed, combusted 47 or 22 mm GelmanA/E filters (nominal pore size = 1.0 pLm).Dry mass of total suspended solids was obtained by drying the filters at 60?Cfor 48 h or until dry mass stabilized. The 22 mm filters were dried as above and then analyzed for stable C and N. Carbonateswere removed from the stable isotope samples by placing filters in a desiccator with 3 mL of concentratedHC1for 4 h. Stream biofilm samples for stable isotope analysis and a mixture of terrestrialleaf litter from the riparian zone of LD1 were also collected at the same time as FPOMsamples. Biofilm samples were scraped with a scalpel from representativelarge gravel or cobble and stored in an acid-washedfilm canister.After drying at 65?Cfor -48 h, leaf litter and biofilm each were ground into fine powders with a mortar and pestle. Stable C and N analyses were done at the University of Alaska FairbanksStable Isotope Facility,using a Europa20-20 continuousflowisotope ratio mass spectrometer.Isotope ratios were expressed in the delta (8) format:813Cor 815N (units of %o) =

(Rsample/Rstandard)

X 1000,

ratio or 15N:14N where R is the 13C:12C ratio,using a bovine protein (peptone)lab standardreferenced against an internationalstandard (N. Haubenstock, University of Alaska Fairbanks, personal communication). Total microbialbiomass was determined using the phospholipid phosphate (PLP)method (Findlay et al. 1989) and microbialcommunity structure was determined using phospholipid fatty acid (PLFA)profiles (Findlay1996) for autumn samples only. Briefly,FPOMsamples collected by centrifugation(9000 x g for 10 min at 4?C)were extractedusing 27.5 mL of a singlebuffphase dichloromethane:methanol:phosphate er (50 mM, pH 7.4) solution (1:2:0.8).After 24 h, 7.5 mL of dichloromethaneand 7.5 mL of deionized water were added, which split the mixture into 2 phases, with the lower, dichloromethane phase containingthe lipid that was recovered and dried. The dried lipid was then resuspended in chloroform and 2 subsamples were removed for determinationof total microbial biomass. These subsamples were placed in glass ampoules, digested to orthophosphate with persulfate,and the phosphate content de-

103

termined colorimetricallyusing a dye-coupled reaction between ammonium molybdate and malachite green. The remainderof the lipid resuspension was partitionedinto neutral,glyco-, and phospholipid using silicic acid column chromatographyand the phospholipidswere recovered. Phospholipidfatty acids were converted into their respective methyl esters by base methanolysis and purified using reverse-phase C18 column chromatography.Purifiedfatty acid methyl esters were identifiedand quantifiedusing gas chromatography.Calculationsof bacterial and eukaryoticC were based on conversion factorsfrom pure and enrichedcultures (Dobbs and Findlay 1993) and published algorithms (Findlayand Dobbs 1993). Relativedigestiveenzymeactivity Ten individuals of E. dilatataand P fasciolaris were collected in the summer at LD1 and BD1. Ten individuals of each species were collected at LD1 in autumn,but 10 individuals of only E. dilatatawere collected at BD1 (no P fasciolaris) because of low abundance.Instead, 10 individuals of P fasciolariswere collected at BD2, -6 km downstream of BD1. Ten individuals of the crayfish Orconectesrusticuswere collected from both BD1 and LD1 in summer and autumn as a controlfor comparisonof mussel digestive enzymes among sites. Digestive fluids from individual mussels were removed by cutting the adductormuscles and then opening the left valve at the hinge. A 25 gauge needle was inserted into the digestive gland area and -100 pL of digestive fluid were removed. For crayfish, the carapace was removed and a 25 gauge needle was used to extract -100 FpLof stomachfluids. Extractedmussel and crayfish digestive fluids were placed into prelabeled1 mL Nunc vials, flash frozen in liquid N, and stored at -80?C until analyzed. Artificialsubstrateswere used to assay activity of 4 classes of digestive enzymes including esterases,lipases, proteases,and carbohydrases. Artificial substrates, obtained from Sigma, St. Louis, Missouri, consisted of methylcoumarin fluorophores (methylumbelliferone [MUF] or methylcoumarinylamide[MCA]),and are covalently bound to different biochemicalmoieties. MUF-P-D-glucosewas used to measure P-glucosidase activity, MUF-palmitatewas used to measure lipase activity,MUF-butyratewas used


104

A. D. CHRISTIAN ET AL.

to measure activity of esterases, and MCA-alanine was used to measure protease activity. Hydrolysis by the digestive enzymes released the fluorophore, and increased fluorescence over time described the relative enzymatic rate for each enzyme class. Enzyme activity was measured as nmol substrate hydrolyzed per min per mL of digestive fluid. This approach has been used successfully to measure digestive enzyme activity of metazoans from both marine and freshwater systems (e.g., Mayer et al. 1997, Smoot and Findlay 2000). Digestive fluid samples were kept at 4?C in the dark and were thawed in an ice bath immediately before enzyme activity analysis; all assays were done at room temperature. Prior to assay, a 20 LLsubsample of digestive fluid was diluted with 6.0 mL of 0.1 M potassium phosphate buffer to obtain a pH of 8.0. This pH level is typical of aquatic invertebrate digestive fluid (Mayer et al. 1997) and was verified by measuring the pH of pooled samples of mussel digestive fluid. The activity of each enzyme class was determined by adding 100 JL of 100 JLMartificial substrate to 900 [LL of diluted digestive fluid, and then continuously measuring (for 12 min) release of fluorophore with a Perkin Elmer LS 50B fluorescence spectrophotometer (excitation 340 nm, emission 450 nm). Quenching of the fluorophore was determined for each artificial substrate by measuring fluorescence of free MUF or free MCA in diluted digestive fluid and then comparing fluorescence of fluorophores in 0.1 M potassium phosphate buffer (pH 8.0). Background fluorescence from autohydrolysis of the artificial substrate was determined by adding substrate to 0.1 M potassium phosphate buffer (pH 8.0). Some substrates are sensitive to autohydrolysis and degrade with repeated freeze-thaw cycles. Thus, aliquots of stock substrate solutions typically were kept in the dark at -70?C until assayed for enzyme activity. Concentration of released fluorophore for each artificial substrate was determined using a serial dilution of free MUF or free MCA in 0.1 M potassium phosphate buffer (pH 8.0). After adjusting for background fluorescence and quenching, the change in fluorescence versus time was converted to molar rates of hydrolysis. Trophic position of each species of mussel and crayfish controls was determined by dividing lipase by protease activity (e.g., Mayer et al. 1997, Smoot and Findlay 2000).

[Volume 23

Musseltissuestableisotopes Following removal of a digestive fluid sample in the field, mussels were measured for total wet mass (shell + tissue) and then were flash frozen. Individuals were brought to the laboratory, thawed, and measured for tissue wet mass and shell wet mass. Tissues were then dried to a stable mass at 60?C (>48 h) to obtain tissue dry mass. After drying, whole soft-tissue samples were ground into a fine powder using a mortar and pestle and then analyzed for stable C and N isotopes, similar to the method described for FPOM.

Statisticalanalysis Average 8 values were used to simulate an integrated food source through time for summer and autumn FPOM for both stable C and N using a 50:50 ratio of summer and autumn FPOM. Stable C isotopes were considered different if their respective ranges did not overlap +2%o (Fry and Sherr 1984, 1989). Stable N isotopes were used to generate estimates of trophic enrichment (TE), which represents the number of trophic levels between an organism and its food base. TE was calculated by subtracting 815N of the food source (865NBAsE)from that of the consumer (S15NcoN) and then dividing the difference by the average enrichment of 3.4%o (range of 2.5-5%o) per trophic level (Minagawa and Wada 1984, McKinney et al. 1999). Microbial community structure of FPOM was determined by analyzing log-transformed mass % values of PLFAs using principal components analysis (PCA) (MiniTabl2, Minitab, State University, Pennsylvania). PCA results were summarized as a scatterplot of factor 1 and factor 2 scores, with samples identified by site. Microbial functional group assignment of fatty acids followed Findlay and Dobbs (1993). A 3-way ANOVA was used to determine if there were species, site, or season differences in activities of each enzyme class. Multiple comparisons were made using the Student's t posthoc test (oL= 0.05) (Jump IN?, version 4, SAS Institute, Inc., Cary, North Carolina). Sequential Bonferroni adjustments were used to maintain an experiment-wise error rate of 0.05 (Lessios 1992).



2004]

105

TABLE1. Mean (?1 SE) concentrationsof seston and its constituentsin stream water.Microbialbiomass is based on phospholipidphosphate (PLP)concentrations,phospholipidfatty acid profiles,and the algorithmsof Findlay and Dobbs (1993) and Dobbs and Findlay (1993).FPOC = fine particulateorganic C, FPON = fine particulateorganicN. BD1 = Big DarbyCreeksite 1, BD2 = Big DarbyCreeksite 2, LD1 = LittleDarbyCreek site.

Site BD1 BD2 LD1

% micro% microMicrobial Micro- % N as bial N Micro- % C as bial C biomass Seston (pmol FPOC bial C micro- as bac- FPON bial N micro- as bacterial terial (mg/L) (mg/L) bial PLP/mL) (mg/L) (mg/L) bial (mg/L) 0.076 82.9 40.6 35.8 25.5 0.063 4.124 (1.682) 25.00(2.29) 1.114 0.399 77.1 41.4 0.046 0.059 128.3 58.6 2.697(0.596) 23.41(0.91) 0.432 0.333 22.9 37.3 86.6 0.025 0.035 140.0 1.249(1.274) 13.62(1.59) 0.264 0.228

Results Seston The amount of seston in stream water from the 3 sites ranged from 1.25 to 4.12 mg/L, with C and N content ranging from 16 to 27% and 1.7 to 2.0%, respectively (Table 1). Living microbial biomass ranged from -15 to 25 pmol PLP/ mL, suggesting that much of the seston C and N was microbial. Bacterial biomass constituted -20 to 60% of the microbial C and N (Table 1). Seston from the Big Darby Creek sites contained significantly higher living biomass than seston

from Little Darby Creek (F = 32.92, df = 2,6, p < 0.001; Table 1). PLFA profiles from seston samples contained 53 different fatty acids. Each site (BD1, BD2, LD1) contained a unique microbial community (Fig. 1). PCA factor 1 separated samples into the 3 sites, with replicate samples grouped by site. Factor 2 separated BD1 from BD2 and LD1. Factors 1 and 2 explained 39.4 and 30.4% of the total variance in the PLFA profiles, respectively. Forty-six individual PLFAs had a component loading >10.51for factor 1 and/or factor 2. Seston from LD1 showed high relative abundances

2.0 -

Microeukarya 1.5-

C\M 0O LL

BD1

1.00.5-

U0.0-

LD1

BD2

Aerobic and -0.5anaerobic bacteria

m mm

-1.0.

1

-I

-1.5

...

I

I

-1.0

0.5

Aerobic and anaerobic bacteria

I 0.0

I 0.5

.

1.0 1.0

1 1.5

Factor 1 Phototrophic microeukarya

FIG. 1. Principalcomponentsanalysis (PCA)of autumn 1998 microbialcommunitystructureat study sites Big Darby 1 (BD1),Big Darby 2 (BD2),and Little Darby 1 (LD1).PCA axes are based on dominantfatty acids within seston.


[Volume 23

A. D. CHRISTIANET AL.

106 14 -

Mussels

Summer FPOM

0 * E. dilatata Z P. fasciolaris

12-

O

Nichols and Gaing 2000 mussels

10-

El

~

FPOM A Terrestrial litter

~[""'"" Nichols and Garling 2000 mussels

8 -

z

Ln

Autumn FPOM --..

~,c

6Autumn FPOM average 4 -

-

/

Nichols and Garling 2000 FPOM

2Nichols and Garling 2000 terrestal lier 0 -33

T -32

litter

Terrestrial

I

I

a

I

I

7

Z

I

-31

-30

-29

-28

-27

-26

-25

-24

03C FIG.2. StableC and N isotopes of the freshwatermussels Elliptiodilatataand Ptychobranchus fasciolaris,fine particulateorganic matter (FPOM),and terrestrialleaf litter from summer (open symbols) and autumn (solid symbols) in the DarbyCreek system. Data from Nichols and Garling(2000)are also shown (shadedsymbols). Ellipses representgroup designations. of the fatty acids 18:3o3, 16:1o9, 16:105, 18:4o3, 16:4ol, 14:0, 18:1w9, and 16:1o13t (ranked by component loading), which commonly occur in phototrophic microeukarya (i.e., algae). Seston from BD2 showed high relative abundances of the fatty acids 16:lo7, brl5:1, 18:1w7, 17:106, i16:0, polyl8:0, a15:0, and i14:0, which, except for polyl8:0, are commonly found in both aerobic and anaerobic bacteria. Seston from BD1 showed high relative abundances of the fatty acids 22:6w3, 20:5o3, 16:4w3, 22:5o3, 20:4w6, 22: 4o6, 20:2, and 10mel8:0, which, except for 10mel8:0, are commonly found in microeukarya. The difference in chain lengths between LD1 and BD1 (16 and 18 carbons vs 20 and 22 carbons) indicated that the 2 sites differed in their microeukarya communities. Observed differences in microbial community structure were consistent with the increasing importance of bacterial biomass to total microbial biomass among the sites, in the order LD1 < BD1 < BD2 (Table 1). Stable isotopeanalysis Terrestrial leaf litter and FPOM during autumn were depleted in '3C (Fig. 2, Table 2) rel-

ative to 13Cwithin stream biofilm. Biofilm was highly enriched in 13C relative to mussel tissue (Table 2) and, thus, was subsequently dismissed as a food source for mussels. Mussels were the most depleted in '3C and the 2 species were more similar to each other within sites than among sites (Fig. 2, Table 2). Leaf litter was depleted in '5N relative to FPOM, which in turn was depleted in 15N relative to biofilm (Table 2). Mussel 8'5N values were similar among species and sites (Fig. 2, Table 2). Species-by-site averages of 815N in mussels were enriched by 2.59 to 2.796%o compared to average summer and autumn FPOM values. These enrichment values translated to TE values of 0.76 to 0.82, indicating a one trophic level difference between the presumed FPOM food resource and mussels. Digestive enzyme analysis Relative activity rates of mussel digestive fluid enzymes were similar among sites within seasons for all enzymes investigated (Table 3). Activity rates in mussels tended to be higher in autumn than late summer, whereas crayfish activity rates varied both seasonally and among sites (Table 3).


107


2004]

TABLE2. Mean (+1 SE) stable C (813C) and N (815N) isotope composition of tissues from the freshwater mussels Elliptiodilatataand Ptychobranchus fasciolarisand their potential food sources.FPOM= fine particulate = organicmatter.BD1 - Big Darby Creek site 1, BD2 = Big Darby Creek site 2, LD1 LittleDarby Creek site.

Source E. dilatata

Season

Site

n

813C

615N

Summer Autumn

BD1 BD1 LD1

8 5 3

-29.2 (0.1) -30.6 (0.3) -31.9 (0.1)

11.5 (0.1) 11.4 (0.2) 11.5(0.1)

-32.1 (0.1)

9.8 (0.2)

-30.4 (0.2) -30.4 (0.2) -31.5 (0.2)

11.1 (0.1) 11.9 (0.1) 11.3 (0.1)

-32.4 (0.2)

9.4 (0.0)

4 1 1 1 3

-29.1 (0.5) -26.6 -26.8 -25.6 -26.3 (0.8)

12.3 (1.6) 6.4 2.5 6.4 5.1 (2.6)

3 3

N&G 2000a

7 3 3

BD1 BD2 LD1

Summer Autumn

P fasciolaris

N&G 2000a

FPOM

Summer Autumn

BD1 BD1 BD2 LD1 Autumn average

48 pIm FPOM

Autumn

N&G 2000a

-30.0 (0.3)

6.2 (0.3)