THE ACTIVITY OF CHTTOBIASE IN THE MEDIUM: A BIOCHEMICAL ESTMATE OF

DEVELOPMENT RATE EN PLANKTONIC CRUSTACEA

A Thesis

Presented to

The Faculty of Graduate Studies

of

The University of Guelph

by

AKASH RENE SASTRI

In partial fulfilment of requirements

for the degree of

Master of Science

April, 2001

O Akash R. Sastri, 200 1

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ABSTRACT

THE ACTIVITY OF CHITOBIASE IN THE MEDIUM: A BIOCHEMICAL

ESTIMATE OF DEVELOPMENT RATE IN PLANKTOMC CRUSTACEA

Akash R. Sastri

University of Guelph, 2001

Advisor:

Professor J-C- Roff

The activity of the molting enzyme chitobiase in the medium surrounding individuals and

populations of planktonic crustacea was investigated. Two applications of this enzyme

assay are presented as methods of estimating development rates in crustacean

zooplankton.

The correspondence between elevated chitobiase activity in the medium and the presence

of exuviae was confirmed in seven fieshwater cladoceran, and one fieshwater and six

marine copepod species. This biochemical cue of the molting event was applied as a

method of estimating the proportion of animals molting in a defined period of time

(development time). An estimate of Daphnia magna (2,000-2,100 p m size class)

development time was in close agreement with that derived by conventional incubations

(70.3 versus 75.1 hours respectively).

Chitobiase activity in the medium was found to Vary with body length in six tteshwater

cladocerans and six marine copepod species. Although the dopes of species specific

regressions differed, a significant common relationship was found (loglo [chitobiase

activity] = -1.19 + 0.89 loglo [body length], r2 = 0.79, p<O.0001). Under steady state

conditions in laboratory cdtures, the rate of decay of chitobiase in the medium was

balanced by its rate of production by molting animais. The rate of decay of the enzyme in

the absence of animais was therefore aIso its rate of production, which is a measure of the

average rate of development of the crustacean zooplankton community. Development

times for a Daphnia magna culture (2 determinations) and a Ceriodaphnia sp. - D.

magna rnixed culture were 65.4 vs. 62 hours, 59 vs. 67 hours and 46.6 vs, 50 hours, as

measured by this application of the chitobiase assay versus conventional molt rate

determinations respectiveIy.

ACKNOWLEDGEMENTS

During the last year of my undergraduate studies, I was exposed to some aspects of aquatic science research that both excited and compelled me to leam more. My sincerest gratitude to my advisor John C. Roff, for facilitating a learning expenence that has far exceeded any possible expectations I might have had when 1 began two years back. I thank John for his guidance, encouragement and infinite patience.

1 wodd also like to thank my advisory cornmittee members, Professor D.H. Lynn, and Professor J.S. Ballantyne for their help with d l manner of questions and carefûl review of my thesis. Some of my field work was conducted in Dorset, Ontario, where Professor N.D. Yan was very kind to provide me facilities, and his thoughtful insights were helpfùl and greatly appreciated.

My lab mates, Warren Cume, Susan Evans, Kem Finlay, Kim Rose and Richard Janutka were together responsible for creating a daily experience in the lab that was always exciting, thought provoking, and above d l fun to participate in, thank you.

I would also like to acknowledge the Department and fellow graduate students, specifically Ken Oakes and the boys in the Ballantyne lab for their fi-iendship and help in the lab. Also my ankle is as good as new, nearly, thanks to the kind efforts of Colin Darling, Michelle Campbell, Susan Evans and James Kowaleski, Cheers!

And lastly, and by no means least, my parents and sister have supported my endeavors in every possible way, for their love and support, 1 am always grateful.

TABLE OF CONTENTS

. ............................................... Chapter 1: Thesis background and rationale Pg 1

1 o d u c o n ................................................................................ Pg- 2

.................... 1.2 Secondary production: Definitions and ecological significance Pg . 3

. 1.2.1 Growth and development rates .............................................. Pg 4

1-32 Conventional methodology and limitations ................................ Pg . 6

1.3 Biochemical methods for estimating growth rates .................................... Pg . 7

. 1.4 Enzymatic indices ......................................................................... Pg 9

. f -5 Chitobiase: Review of previous investigation ........................................ Pg I l

1 .5 . 1 Chitin metabolism: Rationale for study ................................... Pg- 14

1 .52 Chitobiase: Rationale for continued study ................................ Pg- 16

1.6 Thesis outline ............................................................................ Pg . 18

References .................................................................................... Pg- 20

Figure .......................................................................................... Pg . 28

....... . Chapter 2: Chitobiase assay for deterrnining development time in Crustacea Pg 29

Abstract .............................................................................. Pg- 3 O

. .......................................................................... Introduction Pg 32

3 - . Methods and Results ............................................................... Pg 33

Cautions. Optimization and Application ........................................ Pg . 37

References ............................................................................. Pg . 41

Table .................................................................................. Pg- 43

Figure ................................................................................. Pg . 44

Chapter 3: Rate of chitobiase degradation as a measure of development rate in planktonic

....................................................................................... Crustacea Pg . 45

............................................................................... Abstract Pg . 46

Introduction ........................................................................... Pg . 47

Methods ............................................................................... Pg 49 .

............................................................. Results and Calculation Pg . 51

Discussion ............................................................................ Pg 53 .

............................................................................ References Pg . 56

Figures ................................................................................. P g 58

............... Chapter 4: Towards an in silu application of the fiee chitobiase assay Pg . 60

............................................................................... . Abstract Pg 61

.............................................................................. 4.1 Introduction Pg . 62

................................................................................... 4.2 Methods Pg . 62

..................................................................................... 4.3 Results Pg . 64

............................................................................... 4.4 Discussion Pg . 65

..................................................................................... References Pg . 68

.......................................................................................... Figures Pg . 69

Cha~te r 5: Frarnework for in situ applications. sarnpling protocols. modifications . and

.................................................................................... conclusions Pg . 81

. 5.1 Applications. potential limitations. and modifications .............................. Pg 82

5.2 Free ambient chitobiase: What's in your sample? ................................... Pg . 82

5.3 Free ambient chitobiase: Chitobiase discrimination ................................. Pg . 83

5.4 Molting rates: Application and limitations ........................................... Pg . 85

5.5 Molting rates: Handling stress .................... - ..................................... Pg . 86

5.6 Sampling protocol: Thermal layers. population advection, and molting periodicity

. . . ................................... Pg 87 .........................................................

5.7 Sampling protocol : Temperature considerations .................................... Pg . 88

5.8 Isochronal versus non-isochronal development: Potential bias .................... Pg . 88

. 5.9 Microcosm incubations ............................ - ..................................... Pg 89

. 5.10 Modifications: Application of homogenate activities .............................. Pg 91

. 5.1 1 Concluding remarks .............................. - ..................................... Pg 92

. References ...................................................................................... Pg 93

. Figures .......................................................................................... Pg 97

. Appendices ................................................................................... Pg 1 O0

LIST OF FIGURES

C hapter 1.

....... Figure 1. Successive changes in exoskeleton structure during the molt cycle Pg. 28

Chapter 2.

Fig. 1. Relationship between body size and released

non-molting animals ............................. ,,, ...... ., .

chitobiase activity in molted and

.............................. .. Pg. 44

Fig. 1. The relationship between chitobiase activity released by individual animals after

molting and body size in: Ceriodaphnia sp., Daphnia pulex. and Daphnia rnagnrr.

................................................................................................ Pg. 50

Fig. 2. Change of chitobiase activity in whole cultures of Cladocera: Mixed Ceriodaphnia

sp. and Daphnia magna, and Duphnia magna alone. Change of chitobiase activity in

aliquots fkorn cultures fkom which animals have been removed: Mixed Ceriodaphnia sp.

..................................... and Daphnia magna, and Daphnia magna alone .,. Pg. 59

Chapter 4.

Figure 4.1. Cross calibration of fluorescence units (fsu) between Turner Designs (TD

......................... 7000) and Perkin Elmer Luminescence Spectrometer (LS50) Pg. 69

Figure 4.2. Linear regression of chitobiase activity versus body length for Daphnia

magna .......................................................................................... Pg. 70

Figure 4.3. Linear regression of chitobiase activity versus body length for Daphnia pulex

................................................................................................... Pg- 71

Figure 4.4. Linear regression of chitobiase activity versus body length for Ceriodaphnici

sp. ............................................................................................... Pg. 72

Figure 4.5. Linear regression of chitobiase activity versus body length for Daphnia

galeata ......................................................................................... Pg. 73

Figure 4.6. Linear regression of chitobiase activity versus body length for Daphnia

....................................................................................... plicaria. Pg. 74

Figure 4.7. Linear regression of chitobiase activity versus body length for Daphnia

dubia ........................................................................................... Pg- 75

Figure 4.8. Linear regression of chitobiase activity versus length for Holopedium

...................................................................................... gibberum Pg. 76

Figures 4.9.Linear regression of chitobiase activity versus body length for marine

copepods .......................................................................--...--.-..---- Pg- 77

Figure 4.10. Linear regression of chitobiase activity versus body length for al1 species

pooled . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . -. . . . . - -. -. - . -. -. . . .Pg. 78

Figure 4.1 1. T h e coune for of the enzyme-substrate reaction of ambient chitobiase from

seawater. . . . ., . . . -. . . . . . . . . . . . -. . -. . . -. . -. -. . . . . . . . . 79

Figure 4.12. Rate of decay of ambient chitobiase in fieshwater collected fkom Plastic

Lake, Dorset, Ontario . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . - . .Pg. 80

Chapter 5.

Figure 5.1. Rate of ambient chitobiase decay in 0.2 pm-filtered and ~ ~ l t e r e d water

removed from Duphnia magna culture vesse1 in the laboratory . . . . . . . . . . . . ... . . . . . . . .Pg. 97

Fi-we 5.2. Chitobiase activity versus body length in homogenates of non apolytic

Daphnia magna . . . . . . . . . . . . . . . . . . . . . . . . . . - . . . . . *. . . . . . . - -. . . . . . . -. . . . .. . . . -. - - . . . . . . . . . . . Pg. 98

Figure 5.3. Cornparison of chitobiase activity liberated into the medium and resident in

apolytic homogenates of Daphnia magna . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . Pg. 99

LIST OF APPENDICES

Appendix 1 .Composition of synthetic fieshwater .................................... Pg. 1 O 1

Appendix 2. Effect of length of reaction time on hydrolysis of methylumbelliferyl-N-

acetyI-P-D-glucosaminide (MUF-NAG) with medium Erom individual incubations of

molted and non-molted D. magna ................................................... Pg. 102

Appendices 3 a 4 Fluorescence versus tirne for different substrate concentrations of

methylumbelliferyl-N-acte ylglucosamine (MUF-NAG) ...................... Pgs. 1 03 - 1 0 8

Appendix 3g. Linear regression of fluorescence versus methylumbclliferone produced

(fluorescence values = maximal fluorescence from Appendices 3 a to f ) ......... Pg. 1 09

Appendix 4. Time course for decay of chitobiase (unfiltered) released into medium by

individual Daphnia magna ............................................................... Pg. 1 10

Appendix 5. Time course for decay of chitobiase (0.2 pm-filtered) released into medium

by individual Daphnia magna ........................................................... Pg. 1 1 1

Appendix 6. Mean percent chitobiase activity remaining in 0.2 pm-filtered samples of

incubation medium exposed to rnolted Daphnia magna versus tirne (days) .... Pg. 1 12

vii

Appendix 8 . Mean size fiequency distribution of subsamples fiom Daphnia magna

......................................................................... laboratory culture Pg . 1 13

Appendix 9 . Mean size fkequency distribution of subsarnples Daphnia magna .

Ceriodaphnia sp . Iaboratory culture ................................................... Pg . 1 14

Appendix IO . IntermoIt period versus body length . Development times (hours) for

Ceriodaphnia sp.. Duphnia puIex and Daphnia magna ............................. Pg . 1 15

viii

Chapter 1: Thesis background and rationale

1.1 Introduction

The rate o f increase of mass in zooplankton populations is termed secondary production.

The majority of secondary producers in aquatic environments are planktonic crustaceans, 70 %

of which are copepods (Raymont 1980). Owing to their cosmopolitan distribution and great

abundance, planktonic crustacea may represent the dominant muIticellular forrn of animai life on

this planet (Downing 1984)-

Accurate and reproducible estimates of primary and bacterial production exist because of

the development and continued application of radiochernical and biochemical methods to

measure their respective growth rates (Le. Steeman Nielsen 1952, Azam et al. 1983, Berges &

Harrison 1995). By comparison, the scope of secondary production estimates has been limited by

the nature of conventional measures. There have been several unsuccessful attempts to develop

biochemical methods for secondary producers, however, rneasuring rate processes such as

growth in rnulticellular organisms presents several difficulties (see Runge & Roff2000 for

review).

This thesis presents and evaluates two applications of a biochemical method for

estimating development rate in planktonic crustacea. The activity of the molting enzyme,

chitobiase, in the aqueous environment forms the ba i s of this biochemical method. In order to

dernonstrate the value and significance of this new approach, this chapter explores the concepts

of secondary production, particularly the limitations of its conventional measurement, and the

rationale and efficacy of its assessment by alternative biochemical methods, such as the activity

of chitobiase.

1.2 Secondarv Production: Definitions and ecological significance

Production is defined as that arnount of tissue elaborated per unit tirne per unit area (or

per unit volume) regardless of its fate (Downing 1984). This value can be caiculated as the

product of biomass and growth rate, where net growth is the difference between al1 anabolic and

catabolic processes.

Calculation of secondary production is vital to an understanding of material transfer

through aquatic food webs. Knowledge of the efficiency with which carbon, and hence energy,

enters and is dispersed fiom zooplankton comrnunities is important to such practical goals as

management of aquatic resources, assessing the effects of pollution, and an enhanced

understanding of global carbon cycling (Downing 1984).

Runge (1988) explored the question of how and to what extent variation in the physical

environment and phytoplankton production ultimately affect fishery stocks. The principle prey

items for larval fish are zooplankton, the availability of which can have pronounced effects on

the relative success of the early life history of a year class (Lasker 1985; Runge 1988). An

understanding of the factors (both physical and biological) contributing to the relative strength of

this trophic link would be well employed in fisheries management strategies (Vidal 2980;

Downing 1984; Runge 1988; Valiela 1995).

Metabolic rate processes contributing to development in lower trophic levels (Le.,

primary and secondary producers) are responsive to variations in the physical and chernical

environment. For exampIe the changes in measures of metabolic rates of the freshwater

cladoceran, Daphnia magna, are commonly used as an indicator of aquatic toxicity (e-g., Havas

and Likens 1985). Thus, deviations fiom expected production values in these groups might serve

as usefil indices of the effects of pollution at varying spatial and temporal scales.

In terms of the interaction between physical, chemical and biological processes,

knowledge of aquatic production provides an opportunity to examine carbon cycling on a global

scale. The extent to which large aquatic bodies act as sinks, reservoirs, or possible sources of

atmospheric carbon dioxide may contribute to useful general theories about global warming.

Such knowledge should stimulate continued study of those processes, which may act to facilitate

or retard the progression of this phenomenon (e-g. Martin et al. 199 1).

Thus secondary producers serve a potentially important ecological role in aquatic

ecosystems. The extent, to which this potential is presently appreciated, has been limited by the

complexities associated with conventional methods of assessing growth rates in natural

zooplankton populations.

1 -2-1 Secondary production: Growth and development rates

Calculation of production at any particular trophic level requires knowledge of both

biomass and growth rate. The product of these variables defines net production (Downing 1984).

In this context, biomass values are the simpler to determine. Samples are collected at specific

points in time and space, and one of a number of methods used to derive weight (Le., dry weight,

volume of displacement or settling, or length applied to length-weight regressions; Valiela 1995).

Growth rate (g) is calculated as the product of the growth increment (ln(W2'W 1)) and

developrnent time ( I D ) . The growrh increment is the change in mass between two successive

measures of the population's size distribution. This value may be derived by sampling a

progression of specific stages or size classes in the population as a whole in situ. The growth

increments of continuously reproducing populations must be determined in the Iaboratory

through stage or size specific incubations (see Section 1.2.2).

Developmental stages of copepods are defined by gross changes in morphology

associated with a progression through the naupliar and copepodite stages. I n the absence of

stage-specific characteristics (e-g., in cladocerans), a particular stage of development can be set

through the definition of arbitrary size classes. Ii? al1 instances, developmemt time is defined as

the time between successive measurements of mass used to define the grourth increment.

Development tirne defines growth rate (and ultimately production) estimates as rate

processes. This rate-defining component of secondary production estimates is what varies from

one environment to another. Three factors affecting development time and hence growth rate in

zooplankton populations are temperature, food concentration/quality, and body size (Vidal

1980). Huntley and Lopez (1992) discussed a global temperature-based mode1 for growth,

quan t img the over-riding effect of temperature on development. T 'us , im the absence of food

limitation, the relationship between developmental rate and temperature has been used to

estirnate development time (Le., Behlaradek's function; McLaren et al. 7 9 88). Temperature

effects, however, can be complicated by resource limitation as a consequerice of either food

concentration andor the concerted effect of increasing body size (Vidal 1 P80; Berggreen et al.

1988).

The importance of growth and developmental rates as contributors t o production

estimates was demonstrated by cornparison of larvacean to copepod production in tropical waters

(Hopcroft & Roff 1995). The production of Oikopleura dima was comparable to that of the

copepods in the same sarnples, despite a significantly lower biomass- This was attributed to

larvacean growth rates exceeding that of copepods by a factor of ten (Hopcroft & Roff 1995).

A cornplete realization of the impact of growth rates on secondary production estimates

may only be attained through the establishment of a broadly applicable and standardized method.

The need for the development and viability of such a method or methods relies on an

appreciation of the limitations presently imposed on conventional techniques.

1.2.2 Secondary production: Conventional methodolom and limitations

Zooplankton growth rate estimates can be complicated by the nature of a population's

distribution through time. A recognizable progression in the overall size distribution of a

population represents the developrnent of a cohort. The presence of one or more distinct cohorts

can be recognized through a series of temporal samples (Downing 1984). Landry (1978) was

able to distinguish and estimate the growth trajectories of 11 distinct cohorts of Acartia clausi in

this manner. Thus in situ estimates of growth rate c m be obtained where development and

reproduction in a cohort occurs in a synchronous rnanner.

Continuously reproducing populations, however, offer no in sirzr opportunity to calculate

development t h e as there is no recognizable cohort structure (Kirnrnerer 1987). Populations of

this nature exist because females are reproducing throughout the season, resulting in shared

stages of development by individuals fiom different cohorts. Thus, in the absence of distinct

cohorts, growth rate must be detennined through laboratory incubations. The development rate

and growth increments are estirnated by incubating groups sorted by stage or size. Srnaller

anùnals (e-g., copepod nauplii) are difficult to sort by stage, so specific size fiactions are often

incubated. Development of this "artificial cohort" can be followed through time (i.e. Hopcrofr et

al. 1998). The duration of a developmental stage in al1 such incubations can be estimated

through the proportion of anirnals molting during a defined period of time (molt ratio; see

Chapter 2; Chapter 5).

These conventional methods assume exponential growth of the population. The estimated

growth coefficient (g) is applied to the equation; G = eg - 1 to obtain the finite daily growth rate

(G). Production is then calculated as the sum of the product of biomass and growth rate for each

incubated stage or size class (Kirnmerer 1987; Shreeve et al 19%).

While temperature and light c m be adequately controlled, extrapolation of laboratory

estimates to natural populations may be subject to error since a food climate representing in siCu

conditions may be difficult to reproduce (Ikeda & Skjoldal 1980). Thus, conventional assessrnent

of growth rate in crustacean zooplankton populations is cornplex. Designs of secondary

production studies are complicated and time consurning because of extensive sarnpling protocols

and requisite microscopic identification and measurement of thousands of individuals (Huntley

and Lopez 1992; Runge and Roff 2000). Sirnpler and more broadly applicable techniques are

required. The development of alternative methods, such as biochemical estimates of in siru

growth rate, is therefore well justified.

1.3 Biochemical methods for estirnating growth rates

Biochemical methods of estimating growth rates are attractive because they can be simple

and inexpensive to apply. If properly employed, a biochemical method may rapidly facilitate

reproducible estimates of in situ growth rates. Two generd areas of investigation have been

followed with respect to growth rates: changes in both nucleic acid concentrations (i.e.,

RNAiDNA ratios) and the activities of enzymes (Runge and Roff 2000).

Attempts to quanti@ growth in terms of biochemical quantities such as RNAlDNA ratios,

are based on the assumption that DNA concentration (per cell) is constant while RNA

concentration varies with metabolic rate, This assumption does not always hold me. For

example, Sulkin et al. (1975) demonstrated that DNA content per unit biomass decreased with

increasing body mas. DNA content per unit biomass was also found to decrease with increased

growth rate (Ota & Landry 1984). The DNA cornplement has also been found to v q seasonaily

in somatic cells (Brodsky and Ureyvaeva 1985). Both RNA and DNA content are easily

measured using an ethidium bromide fluorometric assay (Le., Karsten and Wollenberg 1 972,

1977). However, this assay is not specific as it measures total RNA and not RNA that only varies

with metabolic rate (RNA and mRNA). This assay of RNA content will therefore include rRNA,

whose variation with growth is more conservative @ u g e & Roff 2000). Thus, even when

present, the relationship between growth rate and RNA/DNA ratios is ofien weak. For example,

RNADNA ratios have been found to be weakly correlated to the growth rate of severai larval

fish species, with most of the variation explained by temperature alone (Buckley 1984).

Furthemore, the relationship between RNA/DNA and growth rate has been found to vary

between closely ielated taxa, and with age within a species (Steinhart & Eckman 1 992; Runge &

RofT2000). As such, it has been suggested that this ratio may be a better descriptor of nutritional

condition rather than growth (Steinhart & Eckrnan 1992, Jones 1 995). Ultimately, however, a

ratio of equivalent quantities is dirnensionless, and as such lack the tirne dimension (Le. T I ) that

defines a rate process. Therefore ratios of biochemical quantities, such as RNA/DNA ratios are

not tnie measures of rate processes such as growth. Thus, a measure that does not quui t ie a rate

process, and which varies within and among species, cannot serve as a routine measure of

growth rate.

Enzymes, however, hold promise as measures of rate processes, such as growth, because

the rates at which they catalyze their specific reactions are expressed in appropriate dimensions

(M-'T'or 'TL) (Runge & Roff 2000). Furthermore, enzyme assays c m be quite simple,

reproducible, and inexpensive. Studies exploring the potential of biochemical quantities (e-g. ,

RNA/DNA ratios, enzyme activities) have not yet developed an accurate measure of growth rate

that is both suitable for routine in situ studies and applicable across species. The conditions

under which the activity of a single biochemical quantity, such as an enzyme, can provide

meaningful estimates of growth rates are discussed in the following section.

1.4 Enzymatic indices

The optimized activity of rate limiting enzymes may be used to measure the rate of flux

of materials through particular metabolic pathways (Newsholrne & Crabtree 1986). This rate of

flux is an accurate measure of growth rate if the enzyme represents the rate lirniting step in a

pathway whose turnover is representative of growth. Therefore the activity of a suitable enzyme

for growth rate estimation, must actually Vary with growth rate, remain independent of similar

sources in other tissues, be representative of growth both within and arnong species, and

represent the particular pathway's rate-limiting step (Berges et al. 1990).

Accompanying an increase in size, is the expectation that the measured activity of an

enzyme associated with a metabolic pool will also increase. Increased enzyme activity with size

should not be mistaken for changes related to growth. The activity of a suitable enzyme must

Vary with growth and not body size alone. This can be resolved by assaying the activity of the

enzyme in animals growing at rates determined by conditions known to modiQ growth (Le.,

temperature, resource concentration).

For exarnple, Bergeron (1990, 1993) proposed a reiationship between aspartate

transcarbamylase (ATC) and mesozooplankton production. However, the relationship between

body size and ATC activity was confounded with body size as Alyse-Danet (1980) observed in

the first investigation of this enzyme. A study of eight enzymes (Berges et aI. 1990) in

homogenates of ArtemiaF.anciscana revealed that only one, nucleoside diphosphate kinase

W P K ) , varied among individuals of the same size growing at different rates and reared under

differing conditions- The remaining seven enzymes were confounded with body size and

variation with growth rates could not be detected-

Continued examination of NDPK (Jones 1994) suggested that the enzyme is not suitable

as a measure of growth rate. The relationship between growth rate and NDPK activity in juvenile

brine shrimp was predictive. However, this relationship differed through adult deveIopmenta.1

stages and NDPK activity itself did not Vary significantly with growth rate in adult stages. Thus,

NDPK activity is not suitable, as it does not vary with growth rate throughout al1 developmental

stages. The NDPK assay may lack specificity because there may be as many as five different

foms (isozyrnes) of the enzyme residing in different tissues (Parks & Aganval 1973; Runge &

Roff 2000). The presence and activity of each of these isozymes may Vary relative to each other

in homogenates of Artemiafianciscana as a h c t i o n of age andor other conditions affecting

growth (Runge & Roff 2000).

Isozymes are variants of an enzyme sharing a specific substrate. However, they are

kinetically different (e.g., Km and V,, values). Any variation in activity at differing

temperakres or locations may be a consequence of adaptive changes in the relative activity of

one isozyme to another. Further, if a relationship between growth rate and enzyme activity exists

for each isozyme, that relationship may Vary between isozymes. Thus, the activity of an enzyme

assayed in whole body homogenates may not be representative of the actud rate of reaction

catalyzed in a tissue of interest. The size range encompassed by most planktonic Crustacea rarely

facilitates enzyme analysis on a specific tissue. Thus, the turnover of a metabolic pool, as

measured by the activity of an enzyme, must be determined in whole body homogenates. The

complexity of relating growth rate to NDPK activity illustrates the problems associated with

such a protocol.

The activity of the chitinolytic enzyme chitobiase has been demonstrated to Vary with

turnover of the crustacean exoskeleton and appears to Vary with growth rate in homogenates

(Buccholz 1 989; Espie & Roff 1 995a, 1 995b). Chitobiase activity, among several zooplankton

species, has also been assayed in the aqueous environment subsequent to molt (Vrba & Macacek

1994; Oosterhuis et al. 2000; Chapters 2, 3, and 4). The presence of chitobiase activity in the

medium (termed 'free7 chitobiase) is significant, because the enzyme c m be assayed free of

interference from potential isozymes of digestive or vesicular origin. This thesis explores the

application of the fkee chitobiase assay as a measure of development time in planktonic

crustacea. The rationale for this approach is based on the previously observed relationships

between chitobiase and development time in homogenates, and the relationships between the

crustacean molt cycle, chitin metabolism, and growth.

1 -5 Chitobiase: Review of previous investigations

Espie and Roff (1 995% 1995b) explored the relationship between chitobiase activity and

growth rate in the cladoceran Daphnia magna. Chitobiase activity in homogenates was first

investigated as a potential measure of the rate of recycling of chitin from the old to new

exoskeleton during the molt cycle (see 1.5.2). The activity of the enzyme was also applied as a

measure of the Fequency of animals in apolysis (an index of development rate) in nahrral

cladoceran populations. The relevant results of these studies are discussed herein to provide

background and rationale for the application of chitobiase activity in this thesis.

Optimizations of homogenization and centrifugation procedures were explored by Espie

and Roff (1 995a, 199%) in an effort to distinguish the various sources of chitobiase (apolytic

from digestive and vesicular) in whole body homogenates of Daphnia magna. In this manner,

they were able to discriminate animals in apolysis (premolt; see 1 S.1, 1.5.2) fiom those in

interrnolt. Peters et al. (1999) recently examined the activity of chitobiase in both the digestive

tract and integurnent of a larger crustacean species, Euphausia stlperba. The size of this

organism did not preclude enzyme analysis on a tissue specific basis. Chitobiase resident in the

digestive tract was found to differ fiom that associated with molting. The enzyme of epiderrnal

origin modulated significantly with the molt cycle, while that resident in the digestive tract did

not.

Based on the activity o f an enzyme, two potential approaches to quanti9 a particular rate

process exist. Espie & Roff s (1945% 1995b) studies of chitobiase and its relationship to growth

examined both. The first approach, that followed by most studies to date, estimated the flux of

chitin fiom the old to new exoskeleton during apolysis (see Section 1 S. 1) as catalyzed by the

optimized activity of chitobiase, in order to quanti@ growth by proportion of tissue (exoskeleton)

to the whole animal (as per Newsholme & Crabtree 1986). Espie and Roff (l995b) found that the

rate of chitin recycling estimated from V,, overestimated the actual rate of chitin synthesis

@off et al. 1994) by approximately 100 foId (2.633 nrnol N-acetyl-P-D-glucosamine (NAG) mg

protein min -' versus the actual rate of 0.0253 m o l NAG mg protein min -'). There are three

potential reasons why chitobiase overestimates the actual rate of chitin flux. 1) The protocol

developed by Espie and Roff (I995a9 1995b) did not adequately discriminate apolytic chitobiase

from digestive sources. 2) Chitobiase is not the rate-lirniting step in the chitin recycling pathway.

3) The optimized conditions used to assay chitobiase are not representative of those in vivo.

The second approach explored by Espie and Roff (l995b) estimated growth in terms of

differences in the relative activity of chitobiase as measured in populations growing at different

rates. An advantage of this approach is that the estimate of development time is not quantified by

the actual rate of the reaction catalyzed by chitobiase. Estimates of development time dependant

on the actual rate of catalysis, must be calculated through the optimized activity of the enzyme

(V,,), a value that may not be representative of the actual in vivo reaction. Further, this rate of

reaction must be converted fkom tissue to whole body rates (Newsholme and Crabtree 1986).

Espie and Roff (1995b) dispensed with these concerns by comparing the total activity of

chitobiase in homogenates of groups of animals with different development times. Chitobiase

activity was found to increase with decreased molt duration. Thus, the greater the fiequency of

animals in apolysis (total increased chitobiase activity), the shorter the duration of the molt cycle.

Since comparison of developrnent rates in one group to another is based on the scale of

chitobiase activity, this is a not a quantitative measure but a relative index of molt duration.

The relationships between chitobiase activity and body size in homogenates of three

different cladoceran species (Daphnia magna, Daphnia galeata, and Daphnia rosea) were found

to be different (Le., diffenng slopes; Espie & Roff 1995b). Therefore, application of this method

to rnixed populations (naturai cornmunities) may prove difficult because a species-specific

calibration would be required. Furthemore, the unavoidable presence of isozymes in whole body

homogenates suggested that calibrations of this sort may be of limited value because the

relationships between different species may also V a r y fiom one spatiaVtempora1 environment to

another.

In summary, discussion of the metabolic role of chitobiase in the chitin-recycling

pathway has been limited to its suitability as a measure of the turnover of this pathway and as an

index of molt duration. These investigations, and those presented in this thesis, are founded on

the relationship between growth and molting in crustaceans. Thus, the following discussion

examines chitin metabolism as it relates to the crustacean molt cycle.

1.5.1 Chitin metabolism: Rationale for s t u d ~

The process by which an arthropod sheds its exoskeleton is referred to as molting. The

series of physiological events that precede and follow the actual molt (ecdysis) are intimately

related to growth. With respect to the potential of chitinolytic enzymes as indices of growth rate,

one must consider the relationship between molting and growth, the arthropod molting cycle

itself, and the structure and composition of the exoskeleton (Figure 1.1).

Metabolizable (assimilated) energy is that proportion of food (energy) consurned that is

devoted to reproduction, growth, maintenance requirements and in the case of crustaceans,

molting (Valiela 1995). Skinner (1985) arbitrarily defines growth as a simple increase in mass. In

crustaceans growth in mass occurs between molts but the rigid exoskeleton imposes a physical

limit to growth in size. Therefore, the only points, in time, at which an increase in size can be

accommodated, are during premolt (proecdysis), ecdysis, and shortly afier molt (early

rnetecdysis). As such, any investigztion into the relationship between growth and the activity of a

chitinolytic enzyme should focus on a cornparison between molt and intermolt. If a measure of a

specific physiological process purports to index growth, then it should likewise illustrate periods

of littie or no growth. To explain this M e r , an understanding of the major events associated

with the molt cycle is necessary (Figure 1.1).

Intermolt is the period of the molt cycle preceding aporysis and following ecdysis. As

such, no chitinolytic activity with respect to molting is observed. The exoskeleton of an

anecdysid animal is cornposed of (fiom the surface Iayer in) a thin non-chitinous epicuticle, a

chitinous procuticle, and a membranous layer resting upon the epidermis (Stevenson 1972).

Apolysis defines the onset of proecdysis (Jenkins & Hinton 1966). Dissolution of the

membranous layer and subsequent separation of the old exoskeleton from the epidermis

characterizes this sub-stage, producing an apolytic space. During apolysis, epidermal cells are

swollen and secrete molting fluid, which includes the chitinolytic enzymes chitinase and

chitobiase, into the apolytic space. Chitinase and chitobiase continue to degrade the postecdysial

layer of the old exoskeleton up to ecdysis itself. Concomitant with this senes of events, the new

epicuticle and preecdysial layers are synthesized pnor to ecdysis. The postecdysial layer of the

new exoskeleton is fully formed during the early metecdysial period (Figure 1.1 ; Skinner 1985).

A large proportion of the crustacean exoskeleton is composed of a polysaccharide of

glucosamine, chitin (Muzzarelli 1977). The significance of this point is twofold: the vast

majority of planktonic secondary producers are in fact crustaceans; and the degradation of the

chitin-protein complex during proecdysis is caused by the hydrolytic action of chitinase and

chitobiase (Muzzarelli 1977). Chitinase hydrolyzes chitin to oligomers and trimers of NAG,

while chitobiase further cleaves to NAG monomers.

At the onset of ecdysis, the exoskeleton begins to split and the animal slowly emerges.

When the exoskeIeton begins to split, the apolytic space becomes continuous with the aqueous

medium. It is then possible to assay the activity of chitobiase, formerly resident in the apolytic

space and now in the externai medium, irnmediately following molt.

1.5.2 Chitobiase: Rationale for continued study

Chitobiase activity has been assayed in both whole body homogenates and in the medium

surrounding molted individuals and populations (Buccholz 1989, Espie & Roff 1995% 1 995b,

Vrba & Macacek 1994, Oosterhuis et al. 2000, Chapters. 2,3, and 4). The activity of both

chitinase and chitobiase has been found to modulate significantly in several species of planktonic

crustacea (Buccholz 1989; Espie & Roff 1995% 199%). Chitobiase activity was found to

increase 10 fold (relative to intermolt levels) during apolysis and r e m to intermolt levels

subsequent to ecdysis in E. superba whiIe a 5 fold increase was observed in Daphnia magna

respectively (Buccholz 1989; Espie & Roff 1995a). Chitobiase activity has also been found to

Vary with growth rate as evidenced by the average activity in populations growing at different

rates (Espie & Roff 1995b).

The assay for chitobiase was refined in the above studies by determiring optimal pH,

temperature, and substrate conditions. Oosterhuis et al. (2000) studied the released chitobiase in

the medium following ecdysis and found the Km value for Temora longicornis to be 55 pmol T',

and in homogenates of Calanoides carinatus, 5 8 ~ 0 1 l 1 -', and Rhincalanus nasîus, 54 prnol * 1'.

These values are comparable to that of Daphnia magna (6 1 .5pmolw 1 -'; Espie & Roff 1995a)

and Daphnia pulicaria (-57 p o l 1 -'; Vrba & hlacacek 1994). Thus, by virtue of kinetic

properties done, chitobiase seems to be fairly conserved among planktonic crustacean species.

Hoppe (1983) examined the efficacy of fluorometric assays using MUF

(methylumbelliferone) substrates to measure the activity of membrane-bound enzymes in water

c o l a microorganisms. Following this methodology, Vrba et al.(1992) investigated the role of

chitobiase as it relates to water coIumn chitin metabolism. They found a significant relationship

between total copepod biomass and the total ambient activity of chitobiase. In order to quantiQ

the relative contribution of crustacean chitobiase to the total pool ofambient chitobiase in the

medium, they exarnined its activity in the medium surrounding individually incubated D.

pulicaria (Vrba & Macacek 1994).

Chitobiase activity in the medium surrounding molted individuals was termed fkee

chitobiase as the other two sources, (Le., bacterial and flagellate) could be removed fkom water

sarnples through selective filtration (as discussed in Chapters 2 , 3 and 4). Vrba and Macacek

(1 994) incubated individuais in small volumes of pond water. In al1 incubations, an elevated

activïty of chitobiase was detected in the medium surrounding molted individuals. This activity

was dways significantly greater than that surrounding non-rnolted individuals. Further, it was

noted that the fiee enzyme remained relatively stable for at least 12 hours, subsequent to which

its activity becarne rather variable. The rate of decay was significantly enhanced by the presence

of microflora. And lastly, perhaps most importantly, the activity of chitobiase released into the

medium was directly proportional to body length of the animals that produced it.

Oosterhuis et al. (2000) examined the relationship between free chitobiase and biomass

production or growth increment, in the marine copepod T. longicornis. They found a

proportionate increase in the activity of fiee chitobiase released by progressively larger stages.

As such, they accurately predicted the change in mass of a synchronous population through an

increase in the overall chitobiase activity in the arnbient medium (corrected for its rate of decay).

Investigations of the relationship(s) between chitobiase Iiberated into the medium by

planktonic crustacea and aspects of their production are limited to two recent studies (Oosterhuis

et al. 2000; Sastri & Roff 2000, Chapter 2). The preliminary results presented by both of these

studies suggest the activity of fiee chitobiase may serve as a powerfid tool in zooplankton

production studies. The establishment of a new method, which is not characterized by the

constraints associated with conventional measures, may accelerate the rate at which useful

secondary production data are accumdated. The ultimate outcome should be an enhanced

understanding of secondary production at greater than local scales.

1.6 Thesis outline

The following chapters of this thesis describe the potential application of a biochemical

cue in the arnbient medium as a method of calculating development time. Chapter 2 examines the

nature of this biochemical cue, in immediate terms such as correspondence to molting and the

different sources and longevity of the enzyme in treated and untreated media. Potential

application of the free chitobiase assay as a method of calculating the proportion of anirnals

molting is proposed.

It has been demonstrated that the activity of chitobiase in homogenates of E. superba, and

D. magna modulates significantiy between intermolt and apolysis (Buchholz 1989, Espie & Roff

1995% 1995b). This increase in total chitobiase activity has been attributed to chitinolytic

activity during apolysis (see Peters et al. 1999 for a more explicit differentiation between

chitobiase associated with molting and that associated with the digestive tract). Thus, the activity

of chitobiase has been demonstrated to Vary with the turnover of chitin in the crustacean

exoskeleton. Chapter 3 of this thesis explores the total turnover of chitobiase in the medium as a

means of calculating development tirne in a laboratory population of D. magna, and a mixed D.

magna-Ceriodaphnia sp. population.

Chapter 4 of this thesis examines the relationship between body size and fiee chitobiase

activity. The relationship is explored with three additional cladoceran species and two marine

copepod species. An overall relationship encompassing al1 species is presented and evaluated.

This chapter also presents some preliminary in siru data on fiee chitobiase in native freshwater

and marine samples. Based on the results presented in Chapters 2,3, and 4, a framework for a

potential in situ application is proposed.

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Stage D 0 to D Stage D2 to D4

Apolytic space E pidermis

Ecdysis

Epicuticle Preecdysial layer Apolytic space Epicuticle Preecdysiai Iayer Epidermis

Stage A Epicuticle Preecdysial layer Epidermis

Stage A to B Stage C Epicuticle Epicuticle Preecysial layer Preecdysial layer

Postecdysial layer Postecdysial layer (fully synthesized)

Epiderrnis Epidermis

Figure 1. I . Successive changes in a crustacean integument during the molt cycle (after Drach 1939; Roff et aI. 1994)

Where stage D O to D 1 represents separation of epidermis fiom exoskeIeton and onset of apolysis; D 2 to D 4, digestion of old postecdysial layer and synthesis of new epicuticle and preecdysial Iayers; ecdysis, molting of old exoskeleton and liberation of molting fluid into the aqueous environment; A-B, de novo synthesis of new postecdysiai Iayer and; C, hIIy synthesized exoskeleton,

Chapter 2: Chitobiase assay for determining development time in

Crustacea

A.R. Sastri and J. C. Roff

To be submitted in Note format to HydPobiologia

Abstract

The proper calculation of secondary production in crustacean zooplankton depends on the

measurement of their growth rates. This in turn requires knowledge of development times and

molting rates. Detexmination of molt rates currently requires prolonged incubations of

individuals or batches of animals, which usually depends on finding the cast exoskeleton

(exuvia), We have found that chitobiase (one of two chitinolytic enzymes), which is released into

the medium at ecdysis (time of molting), serves as a simple and highly accurate method of

determining the proportion of animais molting during a time interval. Presence and activity of

chitobiase is rapidly and easily measured fluorometrically by release of methlyumbelliferone

f?om Methlyumbelliferyl-N-acetyl-glucosamine. The assay requires a single substrate and a short

incubation period of the water in which an animal has resided. Using cultures and natural

populations of fieshwater zooplankton, we determined the validity of this method by establishing

three criteria. 1) Planktonic Crustacea liberate the enzyme chitobiase at molt, as evidenced by

elevated chitobiase activity in the medium surrounding moIted individuals (presence of exuviae)

relative to non-molted individuals. This was established for adults and neonates of Daphnin

magna, neonates of Daphnia pulex, adult Ceriodaphnia sp. and copepodites of freshwater

copepods. 2) Chitobiase activity was measured in individual anirnals as small as 244 pn in

length. 3) The enzyme activity is stable at room temperatures when filtered (0.2 prn).

Developrnent times were calculated from numbers of animals molting as indicated by the

proportion of animals showing elevated chitobiase activity. Developrnent time using the

chitobiase assay was in close agreement with that derived by conventional incubations (70.3

versus 75.1 hours respectively).This method is applicable to ail marine and fieshwater planktonic

crustacea, and eliminates the need for prolonged incubations of animals and the Iabonous

microscopie search for exuviae.

Introduction

Estimation of growth rates in aquatic secondary producers still relies entirely on

conventional techniques. Growth rate (g) is the product of development rate (lm) and the

growth increment. Thus, g = (Ln(W2/WI))iD. While the growth increment (W2/WI ) can be

readify detemiined where developmental stages or size classes are recognized, it is the

developrnent time that is difficuIt and laborious to measure. In the absence of discrete cohorts,

laboratory or field determinations of development time generally require prolonged incubations

of individuals or batches of animals.

To address this difficulty, a biochemical technique has been developed and tested to

rapidly screen dozens or hundreds of individual anirnals in order to determine the fiequency of

rnolting in crustacean zooplankton and thus derive the development time. The method obviates

the need for microscope work and the need to search for cast exuviae. It is based on the enzyme

chitobiase, one of two chitinolytic enzymes found in a diversity of organisms, including al1

crustacea-

Chitin is a simple polymer of P-(1-4) linked N-acetyl-glucosamine and is the primary

structural constituent of al1 arthropod exoskeletons. At apolysis (the start of premolt), the

exoskeleton separates fiom the epidermis. During premolt, the enzymes chitinase and chitobiase

catalyze a partial recycling of chitin from the old to the new exoskeleton. Several studies of

planktonic crustacea have observed increased chitobiase activity during premolt (Buccholz 1989,

Espie & Roff 1995). At the moment of and subsequent to molt (ecdysis), elevated chitobiase

activity can be measured in the medium surrounding the organisms (Vrba & Machacek 1994).

The chitobiase assay is therefore a potential index of actual growth rate (e.g. Espie & Roff 1995,

Oosterhuis et al. 2000).

In another study, we (Sastri and Roff 2000; Chapter 3) showed that the average

development tirne of crustacean zooplankton populations can be derived fiom a knowledge of

the size-specific rate of production of chitobiase and the turnover rate of this enzyme in the

medium. However, this method has not yet been applied to natural populations, and there may be

limitations to its application (see Chapter 4). For example: where development of populations is

not isochronal, where background levels of chitobiase are contributed fiom non-zooplankton

sources or where attention focuses on a particular species, the assay proposed by Sastri and Roff

(2000) may not be appropriate. The objective of the present study was to determine whedier the

activity of chitobiase in the medium following ecdysis could serve as a simple method of

determining the proportion of animals molting during a given time interval, fiom which the

development time cm be derived.

Methods and Results

Individuals were incubated in test tubes containing synthetic fieshwater (see Roff et al.

1994; Appendix 1) for 6 hrs at 22 OC. Volume of medium and enzyme-substrate incubation time

were adjusted depending on the size of animals (Table 2. L), but al1 assays were nin at saturating

substrate concentrations (Appendix 2). A synthetic freshwater was used in order to reduce any

background chitobiase activity. At the conclusion of incubations, the medium was exarnined for

the presence of exuviae, either by eye or under 20 X magnification. Aliquots of 0.7 ml of

medium were removed fiom tubes in which animals had molted, and chitobiase activity was

measured immediately. Chitabiase activity was measured as an increase in fluorescence with

time in 0.7 ml of incubation medium following the addition of 150 pl of 0.4 mm01 (final

concentration) methy lurnbellifery 1 -N-acetyl- p -D-glucosamhide (MUF-NAG) (Sigma C hernical

Co.). Concentrated substrate stock dissolved in Cellosolve (Sigma Chernical Co.) was diluted to

desired concentration in 0.15 M citrate phosphate bufTer, pH 5.5. Medium sample and substrate

were incubated for 10-40 min (see Table 2.1) and the reaction stopped with the addition of 150

p1 of 0.25 N NaOH. Immediately following the addition of NaOH, fluorescence of liberated

rnethylurnbelliferone (MUF) was measured at 360 nm excitation and 450 nm emission using a

Perkin Elmer LS50 Luminescence Spectrometer. Al1 assays were conducted at 22 OC. Controls

(synthetic freshwater) were run (in triplicate) to assess background fluorescence of the substrate

and any background activity associated with the incubation medium itself. Chitobiase activity is

expressed as nrnol MUF liberated per 1 O min (jbr conversion ofjzuorescence values ta chitabiase

activity see Appendices 3a-g).

The reIationship between molting and an elevated chitobiase activity was first examined

using neonates (600-640 pm) of the cladoceran Daphnia pzilex. An asynchronous Iaboratory

culture was used as the source of animals. Cultures were maintained under a 16L:8D hour

photopenod and were fed Scenedesmus sp., ad libitum. Neonates, released by adult females,

were each incubated in 1.5 ml of synthetic freshwater, and exarnined every three hours. The first

individuals to molt were removed and the rernaining individuds were inspected every hou . At

each subsequent interval (each hour), individuals were examined for presence of exuviae and a

0.7-ml aliquot of medium was removed for enzyme assay. We found that the presence of exuviae

was always tied to an elevated chitobiase activity relative to background levels (Table 2.1).

We also tested for the presence of released chitobiase in the medium surrounding other

zooplankton species. assay was applied to adults and neonates of Daphnia magna, neonates

of Ceriadaphnia sp., and copepodites of Diapromus sp. Again, a strict relationship was observed

between production of exuviae and an eIevated chitobiase activity in al1 trials (Table 2.1).

Oosterhuis et al. (2000) have dso used this assay on the marine copepod Temora longicornis.

Given the universality of chitin biochemistry, we believe that this assay is applicable to al1

crustacea.

In order to determine whether the chitobiase assay could measure the proportion of

animals molting, we compared development time calculated by direct observation against that

derived via this chitobiase method. A conventional measure of development time was

determined by incubating individual ( ~ 2 0 ) D. magna (2,000-2,200 pm) in 3 -0-4.0 ml of culture

medium. Each individual was examined every 3 to 6 hours for the presence of exuviae.

DeveIopment time was calculated as twice the mean time to observation of first m o k This value

was not significantly different fiom direct observation (n=10, mean=74.9 , t=-0.0 103, P=0.992)

of the entire intermolt perîod.

To deterrnine the proportion of animais molting using the chitobiase assay, we randomly

removed and incubated 30 anirnals (2,000-2,200 pm) in 3.0 ml of synthetic fieshwater. Animal

incubation periods Iasted 6 hours, at the conclusion of which, aliquots of 0.7 ml were removed

and chitobiase assayed as per above. Sixteen consecutive 6-hou incubations and enzyme assays

were performed on animals initially removed for each interval fiom the same culture vesse1 as

that used for our conventional determination. We randornly chose animals and contulued to

monitor rnolt rate in this manner over 96 hours in order to allow for any possible die1 periodicity

or synchronicity in time of molt in the population. Development times were calculated fiom

numbers of animais molting as indicated by the proportion of animals showing elevated

chitobiase activity. Development tirne using the chitobiase assay was in close agreement with

that derived by conventional incubations, 70.25 (n= 16, S.E.=9.18) versus 75.1 ( ~ 2 0 ,

S.E.=5.75) hours respectively. The incubation medium used for al1 "chitobiase" determinations

of molt rate was synthetic freshwater (Appendix 1). Anba l s were therefore d e d during the

course of each 6 h o u incubation. Thus, we also compared the proportion of animals molting in 6

hour incubations while housed in medium supplemented with food or without ( 1 ~ 1 0 for each

incubation medium).

The duration of an incubation period is dependent, in part, on the length of time

chitobiase in the medium remains detectable at levels above background. Furthemore, a rnethod

which preserves chitobiase activity in samples for long periods may be useful when assessment

of activity is not irnrnediately possible. Thus, we examined the time course for decay of

chitobiase released into the medium following molt (Appendix 4) . Chitobiase activity was

assayed (n=5) at 0, 3,6, 12, and 18 hours. Mean activity remaining at 12 hours was 79.2% of that

measured at time zero (S.E.=5.22), and had declined at 18 hours to 49.5% (S.E.=8.63).

Accordingly, candidate methods to preserve sample activity were examined. These methods

included combinations of fieezing and chernical treatments. Sarnples were initially assayed at

room temperature and again following specific treatments. The best retention of activity, afier 7-

9 days storage at 5 OC , was 85.8% (mean n=10, S.E.= 0.13) of initial activity, following the

addition of 0.1 mm01 DTT (dithiothrietol) and storage at 5 OC. Other suitable treatments included

storage of 0.2 pm-filtered sarnples at -10 and +5 OC. It was found that filtering alone exerted a

major effect on the rate of chitobiase decay in solution. The activity in filtered sarnples (n=l2)

changed little in the fist 48 hours, after which activity decreased but remained consistently

above 60% after 10 days (Appendix 5,6).

Cautions, Optimization and Application

In adult D. magna individuals (2,000-2,200 p), elevated chitobiase activities were

aiways observed in the presence of exuviae. Likewise, the same relationship was observed with

D. pulex neonates (600-640 p), but in one particular tnal, of 10 of 32 individuals molting over

6 hours, chitobiase activity was detected in the medium 1 hour or less before the observation of

exuviae. Vrba & Macacek (1 994) observed a similar phenornenon with Daphnia puliearia. Thus,

if molting rate is assessed simply fiom the presence of chitobiase, an introduction of error (-5%)

might occur. This error was calculated as the product of the proportion of animals molting with

chitobiase activity before evidence of molt (exuviae) to the total number molting (l0/32) and the

reciprocal of the duration of the experiment (1/6). Thus, molt rates cdculated using the

chitobiase method would tend to overestimate the number of animds molting and underestimate

development time compared to methods dependant on an observation of exuviae.

At the conclusion of incubations, care should be taken to confim that no individuals have

died. We found varying degrees of chitobiase activity in the medium surrounding dead

individuals. Variation may depend upon the length of time an individual has been dead, and upon

the extent to which enzyme is liberated from epidermal vesicles or else the digestive tract.

Furthermore, apolytic individuals that die hold the potential to release more enzyme into the

medium following death.

Optimal assessment of chitobiase activity relies on both the nature of the incubation

medium itself and the length of incubation period. If not 0.2 pm-filtered or autoclaved, native

medium rnay also be contaminated with bacterial, flagellate, andior crustacean chitobiase. Thus,

discriminating between background and chitobiase released by smaller individuals may be

difficult. A simple increase in the Iength of enzyme-substrate incubation period may alleviate

d i s , but prolonged enzyme reactions may result in substrate limitation. Thus, in order to

optimize differences in fluorescence between controls and treatments we used pre-filtered

synthetic freshwater or seawater.

However, the use of filtered water precludes feeding during the incubation period,

perhaps artificiaIly proIonging the intermolt period and causing an overestimate of molt period

duration. Thus, the Iength of incubation period was also of import. We found no bias in molt

ratio between animals incubated in synthetic and unfiltered culture medium (t-test, t = -0.234,

P=0.8 18). Regardless of food availability, an apolytic animal is committed to molt (Shreeve et al.

1998). Thus, the length of incubation periods shodd not exceed the duration of the apolytic

phase (i.e.clO% of molt cycle for D. magna reared at 22 OC). Furthemore, the length of an

incubation period is aiso limited by the rate of decay of liberated chitobiase, which is enhanced

by the presence of microorganisrns in the medium (Vrba & Macacek 1994; Oosterhuis et al.

2000; Sastri & Roff 2000). Bacteria may be introduced to the incubation medium, either attached

to the animals' exoskeleton or released fiom the digestive tract. Depending on the species, the

presence of bacteria may also introduce cell-bound chitobiase activity into the incubation

medium. Thus, the longer the incubation period, the greater the potential for an increase in

"background" chitobiase activity. Chitobiase activity assayed in the medium surrounding non-

molted individuals (background) was observed to increase with duration of incubation (Figure

2.1). Therefore, an experimental protocol (duration of incubation etc.) will be a balance between

the duration of apolysis, the total amount of chitobiase produced (crustacean and cell-bound),

and its rate of degradation before assay. The optimd duration for individual D. magna incubated

at 22 "C was found to be 6 hours or less (Figure 2.1).

The incubation protocol should be based on three factors: 1) degree of synchrony or

asynchrony in a population; 2) the rate of degradation of released chitobiase, which is a function

of both temperature and total microbial activity; and the 3) expected duration of the molt cycle (a

function of temperature, food availability and body size (see Vidal 1980, Huntley and Lopez

1992, Hopcrofi and Roff 1995). In an asynchronous population, 30 single animals were

incubated for 6 hours when the expected duration of the moIt cycle was approximately 60 hours.

Thus, during this 6-hou period one wodd expect 3 animals to molt and show chitobiase activity

in the medium. Clearly the more replicates run, the greater the accuracy of the estimate of

development tirne. Many natural populations of planktonic crustacea show some degree of

synchrony in development, including strong die1 patterns of molting (Hopcrofi et al- 1998). This

should be considered in determining population development times. Continuous replicates (as

discussed above) of incubations throughout the course of the expected molt duration will reved

the existence of such die1 cycles.

Sample volumes and incubation times for enzyme-substrate reactions depend on body

size since the amount of enzyme released is a function of body size within a species (Vrba &

Machacek 1994; Oosterhuis et al. 2000). This relationship appears to extend across a number of

species (Fig 2. l), and c m potentially be used as a measure of molt rate in whole crustacean

communities (Sastri and Roff 3000). A divergence between chitobiase activity and background

activity with increased substrate-incubation time was observed (Appendix 2). Thus, greater

sensitivity can be obtained by prolonged enzyme-substrate reaction time. The smallest animals

that were assayed in this manner were Ceriodaphnia sp. neonates (244-304 pm).

This method depends only on detection of the enzyme's activity in sampIes (at a level

significantly above background), not on a measure of the actual rate of reaction. Provided the

incubation and enzyme reaction conditions are optimal (as discussed above), 50% or less of

remaining enzyme activity (relative to the initial reading) will be sufficient to discriminate

molted fkom non-molted individuds. Thus, we investigated the rate of decay of chitobiase

released into the medium and several methods of rnaintaining enzyme lability.

In surnmary, we have shown that the presence of chitobiase in the medium following

ecdysis is a simple surrogate index of molt and hence development time. Development tirne is a

fùnction of temperature, body size, and food concentration (Vidal 1980). The chitobiase method

can be used under any combination of these variables because it is simply an index of the

fiequency of molting animals. Aithough exuviae in Daphnia can be easily seen, those of its

neonates and the smaller stages of copepods are more difficult to find, or may be consumed

following molt. For these reasons, we suspect that there may be substantial biases in some

estimates of molt rates in smaller crustacea. Our method ailows rapid screening of large numbers

of animals and does not require specific calibration. Further, chitobiase activity remains stable

after filtration such that sarnples can be maintained for future deterrnination. The assay is highly

sensitive and can be used on single microcrustacea as small as 244 pm from the piankton and

benthos of both fieshwater and marine environrnents.

References

Buccholz, F. 1989. Molt cycle and seasonal activities of chitinolytic enzymes in the

integument and digestive tract of the Antarctic krill, Euphausia superba. Polar Biol.,

9:3 11-3 17

Espie, P.J., and Roff, J-C. 1995. A biochemical index of duration of the molt cycle for planktonic

Crustacea based on the chitin degrading enzyme, chitobiase. Lirnnol. Oceanogr.,

40: 1028-1034

Hopcroft, R.R., and Roff, J-C. 1995. Zooplankton growth rates: extraordinary production by the

larvacean Oikopleura dioca in tropical waters. J. Plankton Res., 1 7:205-220

HopcrofS R. R., Roff, J. C., Webber, M. K., and Witt, J.D.S. 1998. Zooplankton growth rates:

influence of size and resources in tropical marine copepodites. Mar. Biol., 132: 67-77.

Huntley, M.E., and Lopez, M.D.G. 1992. Temperature-dependant production of marine

copepods: a global synthesis. Amerïcan Naturalist., l4O:2O 1-242

Oosterhuis, S.S., Baas, A.B., and Klein Breteler, W.C.M. 2000. Release of the enzyme

chitobiase by the copepod Ternoru [ongicornis: characteristics and potential tool for

estimating crustacean biomass production in the sea. Mar. Ecol. Prog. Ser., 196: 195-206

Roff, J-C., Kroetsch, J.T., and Clarke, A.J. 1994. A radiochernical method for secondary

production in planktonic cnistacea based on the rate of chitin synthesis. J-Plankton Res-,

l6:96 1-976

Sastri, A.R., and Roff, J-C. 2000. Rate of chitobiase degradation as a measure of development

rate in planktonic crustacea. Can. J. Fish. Aquat. Sci., 57: 1965-1968

Shreeve, R.R., Ward, P., and Murray, A.W.A. 1998. Moulting rates of Calanzis helgolandiczts:

an intercornparison of experimental methods. J. Exp. Mar. Biol. Ecol-, 224: 145-1 54

Vidal, J. 1980. Physioecology of zooplankton. 1, II. Effects of phytopIankton concentration, an4

body size on the growth rate of Calanus pacificus and Pseudocalanus sp. Mar. Biol.,

56: 11 1-134

Vrba, J., and Machacek, J. 1994. ReIease of dissolved extracellular P-N-acetylglucosarninidase

during crustacean molting. Limnol. Oceanogr., 39:7 12-7 16

Table 2.1. Sizes, incubation volumes, and enzyme reaction times for chitobiase detenninations on various species of microcrustaceans-

Species Size (pm) Nurnbers Chitobiase Nurnbers Chitobiase Incubation Substrate not molting activity* molting activity* volume (ml) incubation time

(no exuviac) (exuviae) (min)

Daphnia magna 2700-2900 7 16.3 S.E.=0.825

Daphnia magna 840- 1 050 7 15.9 neonates S.E.=O.S I Daphnia p u k t 600-640 36 12.9 neonates S.E.=0.04 Ceriodaphnia sp. 244-304 17 16-4

S.E.=0.53 Diapromus sp. 600-630 17 17.3

S.E.=0.878

*Chitobiase activity expressed as nrnol methylumbelliferyl (MLTF) liberated during substrate incubation penod. Al1 activity values have been corrected to 3 .O ml incubation volume and for background fluorescence.

O 500 1 O00 1500 2000 2500 3 O00 3 500 4000

Body Iength (pm)

Fig. 2.1. Relationship between body size and released chitobiase activity in molted and non-

rnolting animals. O-Ceriodaphnia sp.; Ei - ~ a ~ h n i a plex; and A-Daphnia magna. Change in background activity after: -6 hours; and . - 12 hours, with non-molted animals.

Chapter 3: Rate of chitobiase degradation as a measure of development rate in planktonic Crustacea

A.R. Sastri and J. C. Roff

Published in October 2000 as a Rapid Communication in

Canadian Journal of Fisheries and Aquutic Sciences 57: 1965- 1968

Abstract:

We have developed a method to determine development tirne (molt rate) in both single and

mked populations of crustacean zooplankton, based on turnover of the chitïnolytic enzyme

chitobiase in the arnbient medium. We examined the relationship between body size and

chitobiase activity released into the medium following molt in three fieshwater cladoceran

species, Ceriodaphnia sp., Daphnia pulex, and Daphnia magna. Chitobiase activity increased

with body length, and a common relationship was observed among al1 three species (r2 = 0.82, p

< 0.0001). Under steady-state conditions in laboratory cultures, the rate of decay of this enzyme

in the medium was balanced by its rate of production by rnolting animals. The rate of decay of

the enzyme in the absence of animals, was therefore also its rate of production, which is a

measure of the average rate of development of the crustacean zooplankton cornrnunity.

Development times for a D. magna culture (2 detemiinations) and a Ceriodaphnia sp. - D.

magna mixed culture were 65.4 vs 62 hours, 59 vs 67 hours, and 46.6 vs 50 hours, as measured

by this 'chitobiase rnethod' versus conventional molt rate determinations respectively.

Introduction

Radiochernical methods for measurement of primary production and bacterial production

in planktonic cornmunities were developed by Steeman-Nielsen (1952) and by Azam et al.

(1 983) respectively, and are now used routinely in field studies. Despite repeated atternpts to

develop similar methods for measuement of growth rates in zooplankton, the field lapsed and

then fell into disrepute following the challenges of Conover and Francis (1973). AIthough a

radiochernicd method to measure growth in zooplankton has been successfully developed (Ro ff

et al- 1994), the method is too complex for routine use. Growth rates in zooplankton are therefore

still generally measured by conventiond incubation techniques, in which a change of body size

or development stage is measured. Growth rate (g) is the product of the reciprocal of

development time (D) and the growth increment, thus: g = (ln(WflI))/D where: W2 and WI

define the growth increment.

In the Crustacea, the dominant group of metazoan zooplankton, the growth incrernent is

readily determined where developmental stages or size classes can be recognized. The major

difficulty associated with measuring growth in crustacean zooplankton is determining

developrnent time, measured as molt rate. This requires incubations of individuals or groups of

animals until molting occurs and cast exoskeletons (exuviae) are produced, or until there is a

sufficient change in size of an artificially created cohort (see e.g. Hopcroft et al. 1998). Whatever

the protocol, such incubations must generally last for several days and involve repeated handling

of animals, with attendant risk of mortality. In addition, exuviae may prove difficult or

impossible to reliably find, especially in smaller animals or carnivorous species.

There have been several attempts to measure zooplankton growth as a fùnction of

biochemical processes or components, typically rates of enzyme activities (EA) or RNA/ DNA

ratios of whole animals. None of these methods has yet become accepted as a standard or widely

applied (see %mge and Roff 2000 for review). Some fundamental problerns include the fact that

EA's are alrnost entirely a function of body size, rather than growth rate (Berges and Ballantyne

1990) both within and among species.

An exception to this rule is the enzyme chitobiase, one of two enzymes that re-cycle

chitin during the molting of d l arthropods. In homogenates of groups of animals from

populations growing at different rates, the activity of chitobiase was correlated with molt rate

(Espie and Roff 1995a). This correlation existed because the titre of chitobiase modulates

strongly during the molt cycle, and its activity within a population is directly related to the

proportion of animals preparing to molt (in pre-molt following apolysis). It is therefore an index

of the frequency of molt (Le. the average development time within a population; Espie and Roff

19950). However, this relationship is species-specific and in homogenates, chitobiase activity in

the apolytic fluid is not distinguished fiom that in the digestive system. Any relationship between

chitobiase activity in homogenates and molt rate would therefore require a re-calibration for each

species (Espie and Roff 1995b). Moreover, such an assay is therefore not applicable to mixed

populations, Le. natural communities.

An increase in chitobiase activity can however be measured in the medium surrounding

an animal, when it is released at the tirne of molt and subsequent to this event (Vrba and

Macacek 1994). This activity of the free enzyme represents only the chitobiase associated with

molting (Vrba and Macacek 1994; Oosterhuis et al. 2000). Within a species, it is strongly related

to the body size of individuals (Vrba and Macacek 1994; Oosterhuis et al. 2000). If the

relationship between released chitobiase activity and body size is sufficiently similar arnong

species within a taxonomic group, then it could provide a direct measure of development time in

nahiral zoopladcion communities. W e explored this possibility by measuring the activity and rate

of decay of the enzyme released from individual afulmais and in laboratory cultures of Cladocera,

as a preliminary step to field trials,

Methods :

Three cladoceran species were chosen because their size ranges overlap: Daphnia magna,

Daphnia pulex, and Ceriodaphnia sp. Al1 cultures, approximately 15 L in volume, of both single

and mixed species, were rnaintained at 22 OC under a 16L:8D hour photoperiod. No attempt was

made to controi the species composition of the food (i-e., mixed fieshwater phytoplankton), or to

keep the cultures fiee of bacteria. In order to obtain an independent measure of development

time, animals from al1 three species corresponding to ten size classes (body length range 250-

3,000 Fm, n = 20-25 per size class) were rnaintained individually in water removed frorn their

specific culture tank. Each individual was inspected every 3-6 hours for up to 96 hours for the

presence of exuviae. Medium was removed and repiaced every 6 hours. Deveïopment time was

calculated as twice the mean time to first rnolt, for ail individuals in a size class. There was no

significant difference between this tirne and the intermolt period observed directly (n=10 for

each size class) between a first and second molt (t-tests, F (-0.68 to 0.28), P=(0.52 to 0.99). The

relationship between body size and development time for al1 three species and al1 size classes

was plotted and described by the following linear regression:

(1) D = 26.45 +- 0.02l(body length) (r2 = 0.91, p < 0.0001; Appendir I O )

Chitobiase assay

Individuals were incubated in test tubes conta i~ng 3.0 mL (D- magna, D. pulex) or 2.0

mL (Ceriodaphnia sp.) of 0.2-pn filtered synthetic fieshwater for 12 hours (D. magna. D.

pulex) and six hours (Ceriodaphnia sp.) at 22 OC. A synthetic freshwater (see Roff et al. 1994;

Appendix I ) , was used to reduce background activity of chitobiase. At the conclusion of

incubations, the medium was examined for the presence of exuviae, either by eye or under 20 X

magnification- Miquots of OJmL of medium were rernoved from tubes in which animals had

molted, and chitobiase activity was measured immediately. Following assay, body length (top of

head to base of tail spine) was measured.

Chitobiase activity was measured as an increase in fluorescence with time in 0.7 mL of

incubation medium following the addition of 150 pL of 0.4 mm01 (final concentration.)

rnethylurnbelliferyl-N-acetyl-P-D-glucosaminide (MUF-NAG from Sigma Chemical Co.)

Concentrated substrate stock dissolved in Cellosolve (Sigma Chemical Co.) was diluted to the

desired concentration in 0.15M citrate phosphate buffer, at pH 5.5. Medium sarnple and substrate

were incubated for 10 minutes (40 minutes for Ceriodaphnia sp.; see Appendix 2 for linearity of

reaction), and the reaction stopped by the addition of 150 pL of 0.25 N NaOH. Following the

addition of NaOH, fluorescence of liberated methylumbelliferyl (MUF) was measured at 360 nm

excitation and 450 nm emission using a Perkin Elmer LS50 Luminescence Spectrometer. Al1

assays were conducted at 22 OC. Controls (synthetic fieshwater) were run (in triplicate) to assess

background fluorescence of the substrate and any background activity associated with the

incubation medium itself. Chitobiase activity was expressed as m o l MUF liberated per 10 min

(Appendix 3a-g). Fluorescence values for molted individuais of Ceriodaphnia sp. were corrected

to incubation volume and enzyme-substrate incubation times used for D. magna and D. pulex

and then converted to chitobiase activity.

Chitobiase activiîy in cultures

Activity of chitobiase over 48 hours was measured every 6 hours in two separate cultures: a

monoculture of D. magna, and a mixed culture of D. magna and Ceriodaphnia sp. Aliquots of

0.7 mL were removed (in triplicate) fiom culture tanks every 6 hours and enzyme activity was

assayed directly as above. The rate of change of activity in 0 . 2 - p filtered aliquots was also

followed over time (i.e. Figure 5.1)

Enzyme activity was also measured in 15.0-mi aliquots removed fiom the culture tanks. Each

aliquot was unfiltered, but inspected to ensure that no cladocera were included, thus preventing

any potential addition of chitobiase over time. The rate of decay was then followed over time.

Resuks and calculation

A single highly significant relationship was found between released chitobiase activity

and body size among the three species (loglo[activity] = -1.75 + 1 .O7 logIo[size], n = 160, r2 =

0.82, p < 0.000 1, Figure 3. l), dthough the regressions are different for each species (see Figures,

4.2-4.4, p < 0.05).

An approximately steady state in chitobiase activity was observed during a 24-hour

period within each culture (Figure 3 2). Therefore, its rate of production (from molting

individuals) must equal its rate of decay (by natural denaturation and decomposition enhanced by

bacterial activity).

In aliquots fiom cultures h m which animals were removed, chitobiase activity decayed

exponentially over 24 hours (Figure. 3.2). Chitobiase activity decayed more slowly in 0.2-pm

filtered water, indicating that the microbial cornrnunity in ouir cultures must be largely

responsible for its decrease.

We can now quanti* the relationship between the rate o f production of chitobiase (due to

molting of anllnds) and its rate of decay (due to the flora in -the cultures) as follows :

(2) (A[CB]/At)tqA = ((C.ni.li.CBi)clD) +/- (A[CB]/At)c

where: [CB] = chitobiase activity; ni = # of anirnals Sn size cIass i; li = mean length of

animais in size class i; CBi = rate of production of chitobiase per animal in size class i; D =

average development time in days (between molts); NA = indicates aliquots fiom which animals

have been removed; c = indicates whole cultures. Note that -under steady state conditions:

(A[CB]/At)c = O.

Three separate assessments of development times were made using this technique, two on

D. magna monocultures and one on a mixed Ceriodaphnia sp. + D. magna culture. The rate of

decay of chitobiase in culture water without animais [(A[CB]/At)NA] was detennined as the total

change in activity over 24 hours, corrected for culture tank v-olume (Figure 3.2). Total chitobiase

production by the populations in each culture [((Cni.li.CBi)cD)] was calculated using the body

size to chitobiase activity regression (Figure 3. l), and animal abundances and size distributions

from three subsarnples (Appendix 7-8) of 250 rnL volume f iom each culture.

There was a close correspondence between development times estimated fiom chitobiase

turnover (eqn. 2) and those derived independently (fiom eqn- 1 ; see Appendix I O ) as follows:

65.4 vs 62 hours, 59 vs 67 hours, and 46.6 vs 50 hours, respectively.

Discussion

Chitobiase activity is known to be a function of body size within a species, both in whole

animal homogenates (Espie and Roff 1995a) and following release after molt (Oosterhuis et al.

2000). We have now shown that chitobiase activity is a function of body size arnong three

cladoceran species, and that a single mass-specific regression describes the relationship. Further,

the rate of decay of this enzyme in the medium can be used as a measure of the average rate of

deveIopment of the crustacean zooplankton community in cultures in the laboratory. The

technique should be applicable to al1 planktonic crustaceans, and to their natural populations in

the field. However, we do not yet know how well a single size-activity regression may describe

relationships in natural communities of zooplankton (see Runge and Roff 2000).

Detennining development tirne in this manner is attractive because of the ease with

which chitobiase is assayed; it is simple, inexpensive, and sensitive. We can measure the enzyme

activity released by a single animal of 244 pm in length. The method also eliminates repeated

handling and sacrifice of individuals, lengthy incubations, and laborious examination for

exuviae. A significant advantage of the method is that in situ water with natural food can be used

as the incubation medium. Our study was conducted at 22 OC. However the method c m be used

at any temperature. A valid calculation of development time requires only that chitobiase activity

in water samples and those used in the construction of chitobiase-body size relationships should

be at the sarne temperature. However, if this is not possible, values can be corrected for

temperature based on knowledge of the enzyme's temperature dependence (see Espie and Roff

1995b).

Chitobiase activity is found naturally in many bodies of water (Vrba and Machacek

1994). Individual animals were therefore incubated in synthetic fieshwater without food in order

to reduce background readings. Note that this does net introduce a bias into the method because

unfed individual animais are only used to establish the size-activity relationship, net to determine

the molt rate itself. Ideally, the incubation periods should be kept to a minimum, in order to

avoid significant decay of the enzyme. Some enzyme activity was detected in non-molting

animds. This is likely due to chitobiase released fiom the digestive tract. Although these values

were low (< 10% of activity in molting animals), we are now exploring the relationship between

the length of incubation period and the background activity.

It should be clearly noted that this method derives an average size-weighted development

time for a population of animais in a given body of water (see eqn. 2 above). The assurnption is

therefore that development times of al1 animais within a container are very similar (referred to as

'isochronality' for developmental stages within a species). However, developrnent is not

isochronal within the cladocera (see eqn. 1 above) or for copepods living under food limited

conditions (e-g. Hopcroft et al. 1998). However, when animals of a restricted size range

dominate the containers, the development times should be accurate and appropriate. Therefore,

this method is potentially applicable to open waters, lakes, and oceans where the size range of

crustacean zooplankton is restricted, or where populations are developing at nearly isochronal

rates.

Most natural water bodies are divided into upper and lower therrnal Iayers containing

smaller and larger crustacean zooplankton respectively. Therefore, it may be possible to apply

this method withh such compartments. Indeed, various strategies might be employed, such as

incubating individuals of a specific size fi-action (e.g. an artificial cohort, see Hopcroft et al.

1998) in rnicrocosms. Then, once the chitobiase activity versus body-size relationship is

developed for the zooplankton of a given region, only the size spectnim and abundance of

animals need be known and the rate of decay of enzyme measured to derive development times.

References

Azarn, F., Fenchel, T., Field, J.G., Gray, J.S., Meyer-Reil, L.A., and Thingstad, F. 1983. The

ecological role of water column microbes in the sea. Mar. EcoI. Prog. Ser., 10: 257-263.

Berges, J. A., and Bailantyne, J. S. 1990. Size-scaiing of whole body maximal enzyme activities

in aquatic crustaceans. Cam J. Fish. Aquat. Sci., 48: 2385-2394.

Conover, R.J., and Francis, V- 1973. The use of radioactive isotopes to measure the transfer of

materials in aquatic food chains. Mar. Biol., 18: 272-283

Espie, P.J., and RoE, J-C. 1995a. A biochemical index of duration of the molt cycle for

planktonic Crustacea based on the chitin degrading enzyme, chitobiase. Limnol.

Oceanogr., 40: 1028-1034.

Espie, P.J., and Roff, J-C. 1995b. Characterization of chitobiase fkom Daphnia magna and its

relation to chitin flux. Physiol. Zool., 68: 727-748.

Hopcrofi, R. R., Roff, J. C., Webber, M. K., and Witt, J.D.S. 1998. Zooplankton growth rates:

influence of size and resources in tropical marine copepodites. Mar. Biol., 132: 67-77.

Oosterhuis, S.S., Baars, M.A., and Klein Breteler, W.C.M. 2000. Release of the enzyme

chitobiase by the copepod Temora longicomis: characteristcs and potential tool for

estimating crustacean biomass production in the sea. Mar. Ecol. Prog. Ser., 196: 1 95-206.

Roff, J-C., Kroetsch, J.T., and Clarke, A.J. 1994. A radiochernical method for secondary

production in planktonic crustacea based on the rate of chitin synthesis. J. Plankton Res,.

16: 961-976.

Runge, J. A., and Roff, J.C. 2000. The measurement of growth and reproductive rates. Ch. 9. In

ICES Zooplankton Methodology Manual. Edited by R.P. Harris, P.H. Wiebe, J. Lenz,

H.R. Skjoldal, and M. Huntley. pp. 401-454.

Steeman-Nielsen, E. 1952. The use of radio-active (c") for measuring organic production in the

sea. J. Cons. Int. Explor. Mer., 18: 11 7-140.

Vrba, J., and Machacek, J. 1994. Release of dissolved extracellular P-N-acetylglucosarninidase

during crustacean molting. LimnoI. Oceanogr., 3 9: 7 12-71 6.

Fig. 3.1. The relationship between chitobiase activity released by individual animals after

molting, and body size in: Ceriodaphnia sp. (O), Daphnia pulex (BI), and Daphnia magna (A) (logio[activity] = -1.75 + 1 .O7 logi&ize], r2 =0.82, p< 0.0001). Each specie's specific regression was significantly different (pc0.05).

Time (hours)

Fig. 3 -2. Change of chitobiase activity in whole cultures of cladocerans: A mixed Ceriodaphnia sp. and Daphnia magna, a Daphnia magna alone. Change of chitobiase activity in aliquots fkom cultures fkom which animals have been removed: A mixed Ceriodaphnia sp. and Daphnia magna, O and 17 Daphnia magna alone.

Chapter 4: Towards an in situ application of the free chitobiase assay for estimating development time in planktonic Crustacea

Abstract

The presence and rate of decay of the chitinolytic e-e, chitobiase, in the medium, has

been used to estimate development t h e in both single and rnixed laboratory populations of

freshwater cladocerans. As a prelirninary investigation into the viability of field applications of

th is approach, it was necessary to determine the extent to which the relationship between

chitobiase activity and body length was conserved arnong several species (both freshwater and

marine) of microcrustacean zooplankton. Samples of both fieshwater and seawater were also

examined in order to determine whether chitobiase (of crustacean origin) could be assayed, and

its rate of decay followed in the natural environment. A measurable rate of chitobiase decay was

observed in native fieshwater sarnples. Furthemore, chitobiase activity, attributed to molting

crustaceans was assayed in both native fieshwater and seawater samples, and distinguished fiom

ce11 bound sources (>0.2 pn). A significant overall regression of chitobiase activity on body size

was observed for 12 fieshwater and marine species; (loglo [chitobiase activity] = - 1.1 9 + 0.89

loglo [length], 2 = 0.79, p<0.000 1). Each of the species specific regressions, however, were

found to be significantly diffèrent from each other and the overai1 regression.

4. I Introduction

Two unique methods of estirnating development time in planktonic crustacea are

described in Chapters 2 and 3. Both methods are based on an assay of chitobiase liberated into

the aqueous environrnent by rnoltïng individuals. Development of these methods in the

laboratory suggest that they hold promise as routine measures of development rate in natural

comunities of planktonic crustacea. A valid in situ application of the chitobiase method (as per

Chapter 3) is dependant on the satisfaction of two conditions. First, estimates of development

rates for natural zooplankton communities (rnixed species populations), require a relationship

between fiee chitobiase activity and body size, that is applicable across species. This relationship

formed the ba i s of the laboratory trials (Chapter 3), but inclusion of other species is fundamental

towards its broader application. Thus, I explored free chitobiase activity in four additional

freshwater cladoceran species and six species of marine copepods. Secondly, chitobiase liberated

by planktonic crustacea must be detectable, and its rate of decay memurable, in the natural

environrnent. Thus, the activity of chitobiase in sarnples of both natural freshwater and marine

environrnents was investigated.

4.2. Methods

The activity of chitobiase liberated by four additional cladoceran species was exarnined

relative to body length. Sampres of HoZopedium gibberurn and Daphnia dubia were collected late

in Septernber, 2000 frorn Plastic and Dickie Lakes, Dorset, Ontario. Chitobiase activity was

assayed within 2 days of sampling. Daphnia galeata and Daphnia pulicariu were collected fiorn

Guelph Lake, Guelph, Ontario, and maintained in the laboratory (as per conditions discussed in

Chapters 2 and 3). Individuais were processed for chitobiase activity within 1 week of collection.

Marine copepods were sampled regularly fkom late July to mid-Ausst 2000 from

Passamaquoddy Bay, New Brunswick. Individuals were incubated within I day of collections.

Free chitobiase was assayed in individual incubations of 2.0-3.0 ml of synthetic

freshwater (Appendix 2 ) for the cladocerans D. galeara, D. dzrbia, D. pukaria, and H

gibberzm. Al1 assays were conducted at 22 OC as per reaction conditions described in Chapters 2

and 3. Marine copepods were individually incubated in 2-04 .O m i synthetic seawater (Crystal

Sea Brand) for 6-9 hours. At the conclusion of incubations, each sample was examined (under 20

X magnification) for the presence of exuviae. Two aliquots of 0.7 ml of incubation medium were

removed for chitobiase assay.

A11 enzyme-substrate reactions were carried out at 14 "C (corresponding to both in situ

and incubation temperatures) with 0.4 mm01 methylurnbelliferyl-N-acetyl-PD-glu cos^

( W - N A G ) dissolved in 0.15 M CPB (pH 5.5). Reactions were stopped after 20 minutes with

the addition of 0.25 N NaOH and I M EDTA. Vrba & Macacek (1 994) attributed cloudiness in

their final solutions to alkalization caused by the addition of NaOH. Clear solutions were only

obtained in seawater with the addition of EDTA. The rate of MUF Iiberation was determined as a

change in fluorescence read on a Turner Designs (TD 7000) fluororneter at 360 nrn excitation

and 450 nm emission. Parallel blanks (synthetic seawater) were nin in triplicate to determine any

background fluorescence. Fluorescence values were cross-calibrated with fluorescence read on a

Perkin Elmer Luminsecence Spectrometer used for al1 fi-eshwater studies (Figue 4.1). In order to

facilitate comparison to fieshwater species, chitobiase activity was corrected to 22 OC, as per the

temperature relationship described by Espie and Roff (1 995a). lmmediately following chitobiase

assays, body length was measured for al1 individuals. H. gibberum was measured as per Yan and

Mackie (1987), D-galeara, D. puliearia and D. dubia as per Chapters 2 and 3 (top of head to base

of tail spine) and the marine copepods length of cephalothorax (as per McLaren et al. 1988).

Aliquots of seawater (n=3 were obtained fiom Passamaquoddy Bay, New Brunswick. Samples

were fust passed through 60-pin mesh to remove any crustaceans. Chitobiase in both 0.2-pm

filtered and unfiltered fractions was assayed in order to distinguish crustacean fkom ceII bound

activity. The linearity of the chitobiase-MUF-NAG (0.4 m o l ) reaction was tested over the

course of 40 minutes in order to confll~n substrate saturation. ALI reactions were stopped with

the addition of 0.25 N NaOH and 1 mol 1-' EDTA.

The rate of decay of chitobiase in sarnples of fieshwater (n=3), collected fiom Plastic

Lake, Ontario was also examined. Samples were initially passed through 60 p.m mesh (to remove

any crustaceans) and the rate of decay monitored. Treatments were assayed three times over 24

hours. Both 0.2-pm filtered and unfiltered sub-sarnples of the unfiltered sarnple were assayed at

each tirne interval. Al1 chitobiase reactions were run with 0.4 mm01 MUF-NAG, for 10 minutes

and stopped with the addition of 0.25 N NaOH.

4.3 Results

Free chitobiase activity versus body length was plotted for a11 species (D. magna, D.

pulex, Ceriodaphnia sp., D. galeatn, D. puticaria. D. dubia, Hgib berum, and marine copepods) .

Each of the species specific regressions (except for K. gibberum) were found to be significant,

p<0.000 1). Al1 species-specific regressions were also found to be significantly different fiom

each other (Figures 4.2-4.9, pc0.005). However, when the data, for al1 species (except H.

gibberurn, see Section 4.4) was pooled, a single significant relationship (logio [chitobiase

activity] = -1.19 +- 0.89 loglo [length], r2 = 0.79, p<0.0001) was found (Figure 4.10). In order to

test the validity of the pooled regression as a measure of any of the specific body Length-

chitobiase relationships, an F-test, pCO.05 was employed. Vaxiation about each individual

regression was compared to that of the overall regression. The F(12,307) = 15.07>2.3, therefore it

c m be concluded that one or more of the species specific regressions are better representative of

variation of each relationship than is the overall regression.

The linear increase of liberated MUF with tirne indicates substrate saturation for both

total (unfiltered) and crustacean (filtered) fractions of seawater (Figure 4.1 1). Cornparison of

filtered and unfiltered fractions suggested significant cell-bound chitobiase activity in the

ambient medium.

A difference in chitobiase activity was also observed between filtered and unfiltered

fieshwater samples, thus indicating significant cell-bound chitobiase activity. The activity of

chitobiase in sub-samples filtered at each time interval, diminished with t h e , while the activity

in the unfiltered fraction did not (Figure 4.12).

4.4 Discussion

The activity of chitobiase released by individual D a p h i a magna, D. pulex, and

Ceriodaphnia sp., followed a similar relationship with body length (Chapter 3). However, the

species-specific regressions were significantly different (Figures 4.1-4.3; p< 0.05). The status of

the additional cladoceran (D. galeata, D. dubia, D. pulicaria and H gibberurn) and copepod

species illustrate a similar condition. The slopes of individual regressions were found to differ

(Figures 4.4-4.9), but the regression through 6 freshwater cladoceran and 6 marine copepod

species examined was significant (Figure 4.10, p<O.001). It was also found that the overall

regression was not as effective in explaining variation of one or more species regressions than

was their own specific regression. Nevertheless, the overall regression was found to be both well

correlated and significant.

The utility o f the cornmon regression is significant in practical ternis. The most accurate

estimate of chitobiase activity liberated by a rnixed population would be obtained using each

species specific regression where each species is equally represented throughout the entire

population size range. An "ideal" population structure such as this, however, is rarely

representative of a natural community. As such, a simpler approach would be to apply the

cornmon relationship according to size, regardless of species composition. Furthemore, the size

ranges al1 of the species included in Figure 4.10 fa11 within the 95% confidence intervals

bounding the common regression. Thus, an estimate of chitobiase activity for a group of species

would only suffer significant error if the size range of a specific group extended beyond the

confidence limits of the common regression.

Investigation of chitobiase activity in other species that encompass the entire body size

range represented in this study, rnay M e r demonstrate the viability of this relationship, with

respect to the application presented in Chapter 3. The observed relationship between chitobiase

activity and body size for H gibberum was not significant and poorly correlated (r2 = 0.10,

Figure 4.8). Thus data for this species was not included in the pooled data set through which the

comrnon regression was applied. Chitobiase activity liberated by molting EL gibberum may be

more appropriateiy applied to another measure of body size such as weight. The exoskeletons the

other species studied are more calcified and IikeIy have a greater chitin content relative to H.

gibberum (personal communication N. Yan). The poor relationship to body length may be due to

the presence of the gel matrix, surrounding this cladoceran, which af5ords it some of the

structural integrity and protection of the more heavily calcified and ngid exoskeletons found in

daphnids and copepods.

The existence of a common relationship for 12 species of planktonic crustacea is

meaningful. Perhaps the most surprising aspect of this finding is that this relationship

encompasses both closely related and relatively disparate species. With respect to the

methodology presented in Chapter 3, a common relationship is important. It is the basis of an

assessment of development time, and hence growth rates in natural mixed populations of

crustacean zooplankton.

Given these initial resuits and the universality of chitobiase activity in crustaceans, it

appears that this relationship may hold for other species. Assuming a taxonomically conserved

fiee chitobiase relationship to body size, an in situ validation of this method depends on whether

the activity of chitobiase (and its rate of decay) in laboratory cultures, is detectable in the natural

environment.

Chitobiase attributed to molting crustaceans was present in samples removed from both

marine and freshwater environrnents. In both instances, activity in the total fraction (Le.

unfiltered) was higher than that in filtered fractions. In a rate of decay experiment, chitobiase

activity in the total fraction of freshwater sarnples remained constant while the activity in the

filtered sub-samples decayed with time (Figure 4.12). Therefore, it is important to distinguish

crustacean chitobiase from cell bound sources. The substrate, MUF-NAG, is not specific to only

the crustacean form of chitobiase. The potential implications of this lack of specificity demands a

discussion of the sources contributing to chitobiase activity in the total fraction (see Chapter 5).

Accordingly, it is important to recognize what the activity in fiitered and unfiltered sarnples

represent and the extent to which discrimination of chitobiase sources is needed.

References

Espie, P.J., and Roff, J-C. 1995. Characterization of chitobiase fiom Daphnia rnagrza and its

relation to chitin f lux Physiol. Zool., 68:727-748

McLaren, I.A., Sevigny, J-M., and Corkett, C.J. 1988. Body sizes, development rates and

genome sizes arnong Calanus species. Hydrobiol., 167/168:275-284

Vrba, J., and Machacek, J. 1994. Release of dissolved extracellular P-N

acetylgIucosarninidase during crustacean molting. Limnol. Oceanogr., 3 9:7 12-7 16

Yan, N.D. and Macke, G.L. 1987. Improved estimation of the dry weight of Holopediurn

gibberurn (Crustacea, Cladocera) using clutch size, a body fat index and lake water total

phosphorus concentration. Cm. J. Fish. Aquat. Sci., 44: 3 8 1-3 89

O 1 O 20 30 40 50 60

Fluorescence (Perkin Elmer LS50 - fsu)

Figure 4.1. Cross calibration of fluorescence units (fsu) between Turner Designs (TD 7000) and Perkin Elmer Luminescence Spectrometer (LS 50). Fluorescence read on both machines at 360 nrn excitation and 450 nm emission. A dilution series of standard chitobiase in 700 pl of 0.2-pm filtered water incubated for 10 minutes with 0.4 mm01 methylumbelliferyl-N-acetyl-P-D- glucosaminide (MUF-NAG).

2.9 3 .O 3.1 3 -3 3 -3 3 -4 3 -5 3 -6

Log Body Length (pm)

Figure 4.2. Linear regression of chitobiase activity versus body length for Daphnia magna. Chitobiase activity is reported as nrnoles methylurnbelliferone (MUF) liberated in 10 minutes from 0.7 ml of medium. Individuals were incubated in 3.0 ml synthetic fkeshwater for 12 h and chitobiase activity assayed in aliquots removed from molted individuals. Where log [chitobiase activity] = -3.654 + 1.6445 log [body length]; r ' = 0.75, n = 95.

2.8 2.9 3.0 3- 1 3 -2 3 -3

Log Body Length (pm)

Figure 4.3. Linear regression of chitobiase activity versus body length for Daphnia pzdex. Chitobiase activity is reported as nrnoles methylurnbelliferone (MUF) liberated in 10 minutes fkom 0.7 ml of medium. hdividuals were incubated in 3.0 ml synthetic fieshwater for 12 h and chitobiase activity assayed in aiiquots removed fiom molted individuais. Where log [chitobiase activity] = -2.86 + 1 -433 log [body length]; r ' = 0.65, n = 52.

2.5 2.6 2.7 2.8 2.9 3 .O

Log Body Length (pm)

Figure 4.4. Linear regression of chitobiase activity versus body len-th for Ceriodaphnia sp. Chitobiase activity is reported as nrnoles methylurnbelliferone (MUF) liberated in 10 minutes fiom 0.7 ml of medium. Individuals were incubated in 3.0 ml synthetic fieshwater for 6 h and chitobiase activity assayed in aliquots removed from molted individuals. Where log [chitobiase activity] = -0.4748 + 06283 log Body length] ; r = O S 1 1, n = 44.

Log Body Length (pm)

Figure 4.5. Linear regression of chitobiase activity versus body length for Daphnia galeata. Chitobiase activity is reported as mo le s methylurnbelliferone (MUF) liberated in 10 minutes fiom 0.7 ml of medium. Individuals were incubated in 3.0 ml synthetic fieshwater for 6 h and chitobiase activity assayed in aliquots removed fiom molted individuals. Where log [chitobiase activity] = -2.26 + 1.29 log [body length]; r =0.60, n = 13.

Log Body Length (pm)

Figure 4.6. Linear regression of chitobiase activity versus body length for Daphnia pulicaria. Chitobiase activity is reported as nrnoles methylumbelliferone (MUF) liberated in 10 minutes fiom 0.7 ml of medium. Individuals were incubated in 3.0 ml synthetic fieshwater for 6 h and chitobiase activity assayed in aliquots removed fiom molted individuals. Where log [chitobiase activity] = -0.62 + 0.71 log pody length]; r =0.85, n = 9.

2.88 2.90 2.92 2.94 2.96 2.98 3.00 3 .O2

Log Body Length (pm)

Figure 4.7. Linear regression of chitobiase activity versus body length for Daphnia dubia . Chitobiase activity is reported as nmoles methylurnbelliferone ( N F ) liberated in 10 minutes fiom 0.7 ml of medium. Individuals were incubated in 3.0 ml synthetic Ereshwater for 6 h and chitobiase activity assayed in aliquots removed fkom molted individuals. Where log [chito biase activity] = -2.735 +1.39 log [body length]; r = 0.724, n= 7.

2.6 2.7 2.8 2.9 3 -0 3.1

Log Body Length (pm)

Figure 4.8. Linear regression of chitobiase activity versus length for Holopediurn gibbertirn . Chitobiase activity is reported as nmoles methylumbelliferone (MUF) Iiberated in 1 0 minutes from 0.7 ml of medium. Individuals were incubated in 3.0 ml synthetic freshwater for 6 h and chitobiase activity assayed in ahquots removed from molted individuals. Where log [chitobiase activity] = 0.69 +O. 167 log pody length]; r =O. 10 1 , n= 20.

Log Body Length (p)

Figures 4.9. Linear regression of chitobiase activity versus body length for marine copepods. Chitobiase activity is reported as mo le s methylurnbelliferone (MUF) liberated in 10 minutes fiom 0.7 ml of medium. lndividuals were incubated in 3.0 ml synthetic seawater for 6 to 9 h and chitobiase activity assayed in aliquots removed fiom molted individuals. Where log [chitobiase activity] =0.848 + 0.1589 log [body length]; r ' =0.458 . total n= 65 ((O), nauplii n= 20, (0) .

Euryternora sp. ( ~ 2 4 ) . Oithona sp. (n=9), Ternora sp. ( ~ 7 ) . Paracalanus sp. (n=2), Centropages sp. (n=2), and Tortanus sp. (n=l))

Log Body Length (pm)

Figure 4.20. Linear regression of chitobiase activity versus body length for dl species pooled (Figures 4.2 -4.9, except 4.8). Chitobiase activity is reported as nrnoles methylumbelliferone (MUF) liberated in 1 O minutes fiom 0.7 ml of medium. Where log [chitobiase activity] = - 1.19 + 0.89 log [body length]; r =0.79, n= 285.

5 10 15 20 25 30 35 40 45

Time (minutes)

Figure 4.1 1. Tirne course for of the enzyme-substrate reaction of ambient chitobiase from seawater. Chitobiase activity expressed as nrnol of MUF liberated per unit time. Reaction was initiated with the addition of 0.4 mm01 MUF-NAG to 0-7 mi of native seawater collected fiom Passamaquoddy Bay, New Brunswick. Bars represent standard error of totd fiaction (n=3) and 0.2 pm filtered &actions (n=3). Linear regression of the total (unfiltered; a) activity versus time = 11.61 + 0.433 r2 =0.97. Linear regression of the crustacean (0 .2 -p filtered; O) activity versus tirne = 9.28 + 0.235 r2 = 0.92.

O 4 8 12 16 20 24

Tirne (hours)

Figure 4.12. Rate of decay of arnbient chitobiase in fieshwater collected fiom Plastic Lake, Dorset, Ontario. Aliquots (n=3) were passed through 60 p mesh and chitobiase activity assayed in 0.7 ml samples. Chitobiase activity is expressed as nmoles MUF liberated in 10 minutes. All reactions were initiated with the addition of 0.4 mm01 MUF-NAG. Samples (0.7 ml) removed fiom aliquots were either 0.2-pm filtered (O) or unfiltered (O) at each time interval. Bars represent standard errors of mean chitobiase activity.

Chapter 5. Framework for in situ applications, sampling protocols,

modifications, and conclusions

5.1 Applications. potential limitations. and modifications

Applications of the chitobiase assay proposed in Chapters 2 and 3 have been discussed in

terms of their feasibility as in situ measures of development tirne (Chapter 4). The following

sections of this chapter discuss the various natural sources of chitobiase in the water colurnn and

to what extent discrimination of these sources is needed. Further, a detailed discussion of molt

rate experiments and the potential for error due to handling stress are presented in order to

demonstrate some advantages and limitations of the method proposed in Chapter 2.

The rernaining sections are concerned with an in situ application of the method presented

in Chapter 3. Thus, an application of such a study requires discussion of preparatory

investigations, sampling protocols, potential biases, and specific modifications of the technique.

5.2. Free ambient chitobiase: What's in your sample?

Chitobiase activity may be of significant ecological importance in the water colwnn. A

number of studies (Le. Hoppe 1983; Vrba et al. 1992; Vrba et a1.1993) have demonstrated the

relative importance of chitin metabolism by a variety of unicellular organisms. There are at Ieast

three known sources of chitobiase resident in the water coIumn. These sources are chitobiase

bound to the ce11 membranes of certain bacterial species, that bound to various flagellates and

ciliates, and that released at ecdysis by crustaceans and other aquatic arthropods.

Cell-bound chitobiase and several other carbohydrate-hydrolyzing enzymes are employed

by some aquatic species of bacteria in DOC and FOC metabolism. Peptidoglycans of microbial

ce11 membranes are in part, composed of N-acetylglucosamine (NAG). Vrba et al. (1993)

demonstrated that the activity of chitobiase bound to the ce11 membranes of the flagellate, Bodo

saltans, and the ciliate, Cyclidium sp., was significantly correlated with total grazing rates on

bacteriai cultures of Aeromonas hydrophila and Alcaligenes xylosoxidans. Al1 three forms of the

enzyme were disthguished kinetically. The bacterial enzyme was of low substrate affinity (Km

>IO0 p o l MUF hour-' 1-'), while both the flagellate and ciliate chitobiase were of high substrate

-1 -1 affmity (Km < 1 pmoI MUF hour 1 ), the crustacean molting enzyme had intermediate Km

values, ranging from 45-65 p o l MUF hotir-' 1-' (Vrba et al. 1993; Vrba & Macacek 1994; Espie

& Roff 1995a ; Vrba et al. 1996; Oosterhuis et al. 2000).

5.3. Free ambient chitobiase: Chitobiase discrimination

Studies of development rate in populations of planktonic Crustacea need not discriminate

chitobiase foms kinetically. Cell-bound chitobiase c m be removed fiom samples using a

0.2 pn-pore filter. A reduction in the overall activity was observed in filtered samples of native

fkeshwater and seawater sarnples reIative to unfiltered samples (Figures 4.1 1 and 4.12). This

difference in chitobiase activity can be attributed to removal of ce11 bound activity by filtering.

However, the removal of unicellular organisms and ce11 bound chitobiase can only be inferred by

the reduced rate of chitobiase decay in filtered versus unfiltered samples in laboratory

experiments (Figure 5.1).

Filtering should always be conducted gently in order to avoid shearing of cell-bound

chitobiase fiom rnicroorganisms residing in native samples (Oosterhuis et al. 2000). Assessing a

rate of decay based on the presence of both crustacean and unicellular chitobiase may not be

suitable for estimates of development time. Unicellular chitobiase sheared from cell membranes

will overestimate total activity and bias a rate of decay measurement.

The total fraction of chitobiase was observed to decrease in samples fkee of crustaceans

(Oosterhuis et al. 2000; Figure 3.2). However, a measurable decay rate was not observed in the

total fraction of samples fiom Plastic Lake (Figure 4.12). The rate of free chitobiase (filtered)

decay, however, was measurable. These observations (Figure 4.12) suggest that samples

removed from the environrnent should be initially 0.2 pm-filtered to determine the relative

contribution of chitobiase by molting crustaceans. If ce11 bound activity is high relative to

putative crustacean sources, then both 0 . 2 - p filtered and unfiltered sub-samples of aliquots

removed to monitor the rate of decay of chitobiase should also be m. Thus, filtering sub-

samples removed penodically from an unfiltered aliquot should represent the arnbient rate of fiee

chitobiase decay.

If ce11 bound activity is low relative to crustacean activity, a filtering protocol may not be

necessary. For instance, there was little or no difference in the initial activity of 0.2-pm filtered

and unfiltered samples removed from cladoceran laboratory cultures (Figure 3.2 and 5.1).

Indicating that perhaps microorganisms resident in the cladoceran laboratory cultures have little

or no cell-bound chitobiase activity. This observation is in agreement with Vrba et al. (1992)

who found that chitobiase activity is not universal to al1 bacterial species (e.g., Akaligenes

xylosoxidms). The presence of microorganisms in unfîltered samples was therefore inferred by

enhanced rates of chitobiase decay relative to filtered samples (Figure 5.1). Alternatively,

chitobiase of microbial origin may have been overwhelmed by the cladoceran contributions in

the limited volume of the laboratory culture vessel.

In summary, the presence of chitobiase and its rate of decay in natural aquatic

environments was measurable. Discrimination of crustacean chitobiase may be accomplished,

where necessary, through a filtering protocol. Further, results presented in this thesis suggest that

a cornmon chitobiase body size relationship may exist. Thus, an estimate of the development rate

of naturai communities of planktonic crustacea may be possible. The following sections explore

aspects of a sampling protocol, limitations and associated biases, as weli as potentid

modifications of the presented methods in order to establish a k e w o r k for application of the

chitobiase method for routine in situ studies.

5.4 Molting rates: Application and limitations

In situ estimates of growth rate in continuously reproducing populations are difficult

because cohorts are not distinguishable; there is therefore no recognizable progression in the

relative size of stagekize classes of a population. Thus, we must resort to incubations of animals,

either sorted for stage or as an artificial cohort (Tranter 1976; Kimrnerer & McKinnon 1987).

Production, in this instance, is calculated as the product of biomass and growth rate surnrned for

d l sizektage classes (Kirnrnerer 1987).

Stage duration can be determined through molting experiments, where the reciprocal of

molt rate equals development time. Molting rates of specific size classes or stages may need to

be estimated for several types of studies including production in continuously reproducing

populations, and body size/resource concentration studies (e.g. Berggreen et al. 1988; Peterson et

al. 1991 ; Hopcroft & Roff 1995; Shreeve et al. 1998)

Ail conventional molting rate tech~ques (see Shreeve et al. 1998 for review) require an

estimate of the nurnber of animals molting in a defined period of time. This value c m be

estimated as the ratio of exuviae to the total number of individuals or the proportion of animals

progressing to the next stage of development.

A vaiid estimate of stage duration is contingent on the satisfaction of two assumptions:

that the age of individuals within an incubated size class is uniform; and that molting behavior

has not been compromised by capture and handling (Runge et al. 1985). The chitobiase assay

(Chapter 2) is a method that, like conventional methods, relies on uniformity of age within the

incubated size class. M e n the age distribution within isolated size classes fiom natural

popdations is not completeIy uniform, replicate incubations should be conducted (as per Chapter

2)-

5.5 Molting; rates: Handlïng; stress

Several studies have observed deviations fiom normal molting behavior as a consequence

of capture and handling (Miller et aI. 1984; Runge et al. 1985). There is some debate as to what

extent handling affects molting. In some instances, moIt rate was thought to have been artificially

elevated as a consequence of handling (e.g. Miller et al. 1984). Runge et al. (1985) reported that

during some incubations of Calanusfinmarchicus and Calmus glacialis, individuals with

damaged exoskeletons were not able to molt completely. Although injured individuah were not

able to complete ecdysis, the initiation of molting (apolysis) was not influenced. Provided the

time at which animals enter ecdysis remains unaf5ected by handling, a stress of this nature should

not bias molt rate estimates using the chitobiase method. As observed with D. pulex neonates,

chitobiase can be assayed in the medium before complete shedding of the exoskeleton (Chapter

2). Therefore, a complete molt is not required. Shreeve et al. (1998) compared four different

methods of estimating molt rate. No significant difference in molt rates was observed between

any of the methods regardless of the extent to which animals were handled. Regardless of the

incubation protocol and the extent of handling required, any manipulation (i.e. sorting with

pipettes andor filtering through mesh) should be conducted carefidly with attention to "gentle"

techniques (e-g- Peterson et ai- 199 1).

5.6 Sarn~ling Protocol: Thermai layers. ~ o ~ u l a t i o n advection and mol t in~ ~e r iod i c i t~

Chitobiase activity assayed in the environrnent should be ascribed to only those

individuals sampled. Sources of chitobiase released by organisms outside of the study population

will tend to overestimate the mean development time. Thus, a suitable sampling protocol shouid

consider the presence of organisms and their molting behavior in both spatial and temporal

terms,

Smaller animals of a similar size range tend to be restricted to thermal layers in stratified

bodies of water, while larger anirnals are typically found outside this range (Roff, personal

communication). Development time of a population residing within the upper thermal layer can

be estimated accurately, because of a narrow size range and limited opportunity for input of

chitobiase fiom outside sources. However, if the water column is not completely stratified, there

may be potential for chitobiase input frorn other sources.

In completely mixed environments, ambient chitobiase activity should be consistent

throughout the range of the population. In marine environments, the periodic advection of

organisms (i-e. due to strong tidal patterns) may have to be addressed. Thus, sampling of water

for chitobiase assays must be limited in time and space to where the target populations reside. In

this contexq, serial sampling of the water column should identifi if there is any molting

periodicity and strong vertical migratory pattems.

5.7 Sarnpling protocol : Temperature considerations

Chitobiase activity in water sarnples and those used in the construction of chitobiase-

body size relationships must be compared at the same temperature. However, if this is not

possible, values can be corrected for temperature based on knowledge of the enzyme's

temperature dependence (see Buccholz 1989; Espie and Roff 1995a).

Determination of the rate of decay of chitobiase is necessary for the calculation

chitobiase turnover (Equation 2, Chapter 3). Temperature exerts an effect on the rate of decay of

chitobiase (Oosterhuis et al. 2000), where a temperature increase, increases the rate of decay of

Iiberated chitobiase. Therefore, if strong temporal and spatial molting patterns are observed, the

specific temperature regime associated with these patterns should be identified, and chitobiase

activity assayed accordingly.

5.8 Isochronal versus non-isochronal development: Potential bias

Development time estimates are average size-weighted values for the population (see

Equation 2 Chapter 3). Correct interpretations of estimates are based on the degree of population

isochronality, if the size fiequency distribution then exerts a bias, and whether or not incubations

of specific groups are required Development time in planktonic crustacea c m proceed

isochronally or non-isochronally. In populations of food-satiated copepods, development time is

isochronal (Le. development time remains constant throughout ail life history stages; Berggreen

et al. 1988; McLaren et al. 1988). For food-limited copepods and cladocerans, development time

increases with body size (Hopcroft et al. 1998, Chapter 2,3).

In an isochronal population, stage duration remains constant with size. Therefore an

estimate of development time based on the turnover of chitobiase is representative of the

population as a whole. The total chitobiase contribution is calculated as the sum of chitobiase

conûibuted by al1 individuals in al1 size classes. This total contribution is uniformly distributed

throughout the population and an estimate for an "average-sized animal" is derived. Thus, the

estimate of stage duration is not biased by the size distribution of the population because the

development time of an average mimal is thc same as any other animal regardless of size.

Increasing stage duration with size however can bias this method. Since the estimate is an

average size weighted value, it is assurned that al1 individuals share the sarne or simila.

development times. However, when development time increases with size, the caiculation of

mean development time is only useful as a measure of growth rate in populations of a restncted

size range or in which the number of individuals represented in each size class is equal (Le., a

well established continuously reproducing population where the rate of recruitment equals

mortality rate).

A bias rnay exist in a popuIation with one or more size cIasses dominating the population.

Dependant on the size range of the population, the average development time estimated by the

chitobiase method would overestimate development time if the mode of the distribution were

skewed to the left (Le. larger size classes). Likewise an underestirnate would be calculated when

the population was skewed to the right.

5.9 Microcosm incubations

An estirnate of the average development tirne of natural copepod populations that is

greater thm the stage duration predicted by temperature (Le. Behlaradek's function; McLaren et

al.1988; Huntley & Lopez 1992) suggests food limitation and a departure fiom isochronal

development. If, as in the laboratory populations of cladocerans (Chapter 3), the size range is

limited, then estimates wiU be meaningfül. However, when the size range of a non-isochronal

population precludes an accurate estimate, one should consider incubations of specific size

fiactions (Le. Hopcroft & Roff 1995).

Estimates of the mean development time of size fractions should always employ gentle

size fiactionating techniques so as not to affect molting behavior (i.e. Runge et al. 1985; Peterson

et al. 199 1). Most importantly, however, estimates must allow for the presence of chitobiase

fiom al1 in siîu sources, and maintain the in situ temperature regime. The rate of decay of

ambient chitobiase is mediated in part by the presence of the native microflora. An accurate

reflection of the rate of decay requires that housed anha l s be maintained at in situ temperatures-

Furthermore, employing native water sources for incubations best approximates the short-term in

situ food climate.

The length of a microcosm incubation is determined by the rate of decay of chitobiase

initially present the medium. If native water is used for microcosm incubations, then chitobiase

contributions fiom the entire population will initially be present. An estirnate based on the initial

rate of decay will overestimate the development time of an incubated size fraction. Continuous

monitoring of chitobiase activity should reveal a general decrease in ambient activity up to that

point where activity is representative of the incubated size fraction only. This is the point where

contributions of chitobiase by the incubated animals are balanced by their decay. A subsequent

increase in activity will be a consequence of an overall progression in the size of the incubated

size fiaction (i.e., Oosterhuis et al. 2000). Any deviations fiom a steady state can then be

accomrnodated as per Equation 2, Chapter 3.

5.10 Modifications: Application of homogenate activity to chitobiase-body size relationship

The construction of a body size versus a fiee chitobiase activity relationship is necessary

for estimates of development tixne employing the method proposed in Chapter 3. Chitobiase

activity is assayed in the medium surrounding individuals that have molted. The duration of such

incubations should be kept to a minimum (see Chapter 2) in order to avoid significant decay of

the enzyrne before assay. Thus, in an effort to avoid this potential for error, 1 explored the

activity of chitobiase in individual homogenates of Daphnicz magna (as per Espie & Roff 1995%

1995b).

Animais were first inspected under the microscope (40X) for the presence of an apolytic

space at the distal end of the tailspine. Each apolytic animal was hand homogenized in 3.0. ml of

0.15 M citrate phosphate buffer + sucrose solution. Chitobiase activity was assayed as per

Chapters 2 and 3 with the addition of 0.4 mm01 MUF-NAG to a 0.7 ml aliquot of homogenate.

Al1 reactions were stopped after 10 min with the addition of 0.25 N NaOH. The activity of the

enzyme in homogenates of non-apolytic animals was also assayed, and activity versus body size

plotted (Figure 5.2). The size of each apolytic animal was recorded and chitobiase activity of an

equivalent non-apolytic animal calculated fiom the relationship of Figure 5.2. Thus, the

difference in chitobiase activity in apolytic an non-apolytic animals was calculated by

subtraction and plotted against chitobiase activity liberated into the medium by Daphnia magna

(Figure 5.3).

The difference between chitobiase activity in apolytic and non-apolytic homogenates

appears to be comparable to that of the enzyme liberated into the medium by molted animals.

This activity is variable relative to that liberated into the medium. However, assaying chitobiase

activity in homogenates may be more accurate because the potential for decay of the liberated

enzyme is eliminated. The variability of chitobiase activity in homogenates may be related to

length of time the animal has been apolytic. Thus, animals assayed early in apolysis may have

lower chitobiase activities relative to those assayed irnrnediately before ecdysis.

5.1 1 Concludina remarks

The preceding discussion emphasizes specific details of how both of the methods

presented in this thesis are best applied to routine application. Where possible some of these

details have been illustrated with preliminary data on chitobiase activity in both natural marine

and fieshwater samples. As stated on several occasions throughout the body of this thesis, these

methods were developed for routine in siîu application. Both laboratory results and preliminary

data suggest that if properly applied, these methods will facilitate rapid and accurate assessments

of zooplankton development rates

References

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copepod Acartia tonsa durhg development: implications for determination of copepod

production. Mar. Biol., 99: 34 1-352

Buccholz, F. 1989. Molt cycle and seasonal activities of chitinolytic enzymes in the

integurnent and digestive tract of the Antarctic krill. Euphausia superbn. Polar Biol.,

9:3ll-317

Espie, P.J., and RoE, J-C. 1995. A biochernical index of duration of the molt cycle for

planktonic Crustacea based on the chitin degrading enzyme, chitobiase. Lirnnol.

Oceanogr., 40: 1028-1034

Espie, P.J., and Roff, J-C. 1995. Characterization of chitobiase from Daphnia magna and its

relation to chitin flux. Physiol. Zool., 68:727-748

Hopcroft, R.R., and Roff, J-C. 1995. Zooplankton growth rates: extraordinary production by the

larvacean Oikopleura dioca in tropical waters. J. Plankton Res., 17:205-220

Hopcroft, R. R., Roff, J. C., Webber? M. K., and Witt, J.D.S. 1998. Zooplankton growth rates:

influence of size and resources in tropical marîne copepodites. Mar.Biol., 132: 67-77

Hoppe, H-G. 1983. Significance of exoenzymatic activities in the ecology of brackish water:

measurements by means of methylubelliferyl-substrates. Mar. Ecol. Prog. Ser., 1 1 :299

308

Huntley, M.E., and Lopez, M.D.G. 1992. Temperature-dependant production of marine

copepods: a global synthesis. American Naturalist., 140:20 1-242

Kirnmerer, W.J. 1987. The theory of secondary production calculations for continuously

reproducing populations. Limnol. Oceanogr., 32: 1- 13

Kirnrnerer, W.J. and McKinnon, A.D. 1987. Growth, rnortality and secondary production of the

copepod Acarfia tranteri in Westernport Bay, Australia. Limnol. Oceanogr., 32: 14-28

McLaren, I.A., Sevigny, J-M., and Corkett, C.J. 1988. Body sizes, development rates and

genome sizes among Calanus species. Hydrobiol., 167/168:275-284

Miller, C.B., Frost, B. W., Batchelder, H-P, Clemons, M.J., and Conway, R.E. 1984. Life

histories of large, grazing copepods in a subarctic gyre: Neocalanus plurnchrus,

Neocalanus cristasus, and Eucalanus bungii in the Northeast Pacific. Prog

Oceanography., 13 :20 1-243

Oosterhuis, S.S., Baars, A.B., and Klein Breteler, W.C.M. 2000. Release of the enzyme

chitobiase by the copepod Temora langicornis: characteristics and potential tool for

estimating crustacean biomass production in the sea. Mar. Ecol. Prog. Ser., 196: 195-206

Peterson, W.T., Tiselius, P., and Kimboe, T. 199 1. Copepod egg production, moulting and

growth rates, and secondary production, in the Skagerrak in August 1988. J. Plankton

Res., 13:131-154

Runge, J.A., McLaren, I.A., Corkett, C.J. and Koslow, J.A. 1985. Molting rates and cohort

development of CalanusjiPlmarchicus and Cu~unus glacialis in the sea off southwest

Nova Scotia. Mar. Biol., 86:241-246

Shreeve, R.R., Ward, P., and Murray, A.W.A. 1998. Moulting rates of Calanz~~ helgolundicus:

an intercornparison of experirnental methods. J. Exp. Mar. BioLEcol., 224: 145-154

Tranter, D.J. 1976. Herbivore production. In Ecology of seas. Pp. 186-224. D.H. Cushing and J.J.

Walsh (eds.). W .B. Saunders. Philadelphia

Vrba, J., Nedoma, J., Simek, K., and Seda, J. 1992. Microbial decomposition of polymer organic

matter related to plankton development in a reservoir: activity of a-, P-glucosidase, and

P-N-acetylglucosaminidase and uptake of N-acetylglucosamine. Archiv. F.

Hydrobiologie., 126: 193-21 1

Vrba, J., Simek, K., Nedoma, J,, and Hartman, P. 1993.4-methylumbellifery-PN

acetylglucosarninide hydrolysis by a high affinïty enzyme, a putative marker of protozoan

bacterivory. Appl. Environ. Microbiol., SWO9 1-3 10 1

Vrba, J., and Machacek, J. 1994- Release of dissolved extracellular P-N

acetylglucosaminidase during crustacean molting. Limnol. Oceanogr., 3 9:7 1 2-7 16

Vrba, J.,. Simek, K., Pemthaler, J., and Psemer, R. 1996. Evaluation of extracellular, high

affînity B-N-acetylglucosaminidase measurements fiom Ereshwater lakes: An enzyme

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Figure 5.1. Rate of arnbient chitobiase decay in 0.2 pn filtered and unfiltered water removed fiom Daphnia magna culture vesse1 in the laboratory. Aliquots were removed at tîme O and passed through a 60 pm mesh to remove animals. Aliquots were either filtered (O; n=3) or unfiltered (a; ~ 3 ) . Chitobiase activity was assayed in 0.7 ml sarnples every 3 to 6 hours from reactions with 0.4 rnrnol MUF-NAG for 1 O minutes. Bars represent standard errors of mean chitobiase activity.

500 1 O00 1500 2000 2500 3000 3500 4000

Body LengSi (pm)

Figure 5.2. Chitobiase activity in non-apolytic homogenates of Duphnia magna (n=16). Activity is expressed as nmoles methylurnbelliferyl (MUF) liberated in 10 minutes. Animals were homogenized in 3.0 ml of citrate-phosphate buffer, pH 5.5. Fluorescence read at 360 nm excitation and 450 nm emission on Perkin Elmer (LS 50) Spectrorneter: regression of activity versus body length (pm); Activity = -1 39.33 + 0.754 (body length), 8 = 0.75, p<0.000 1.

500 1 O00 1500 2000 2500 3 O00 3500 4000.

Body Length (pm)

Figure 5.3. A cornparison of chitobiase activity (nmoles methylurnbelliferyl (MUF) liberated in 10 min) liberated into the medium (O) and resident in the apolytic space ( a ) of homogenates of Daphnia magna.

Appendices.

Appendix 1 Synthetic fieshwater (as per Roff et al. 1994). AU reagents diluted in Nanopure filtered water. Bracketed values indicate composition of synthetic freshwater used for incubations of Daphnia magna.

NaHC03 0.048 g (0.096 g). 1 "

Cas04 .2Hz0 0.038 g (0.076 g). 1 *' MgSO4.7H20 0.061 5g (O. 123 g)- 1 -' KCl 0.0005g (0.001 g)- 1 -'

5 1 O 15 20 25 30 35 40 45

Time (minutes)

Appendix 2. Effect of length of reaction time on hydrolysis of methylumbelliferyl-N-acetyl-PD- glucosaminide (MUF-NAG) with 0.7 ml of medium fkom individual Daphnia magna (2,000- 2,100 pm) incubated for 6 h (n = 7 moIted (O), versus n = 7 non-molted (O)). Linear regressions of chitobiase activity (nrnol MUF liberated) versus tirne are:

= Molted, a = 4.013, b = 24.964, r2 = 0.8142, p < 0.0001. O =Non-molted, a = 0.279, b= 13.831, r2 = 0.335, n.s. Bars represent standard errors of mean chitobiase activity of molted and non molted individuals.

O 1 O 20 30 40 50 60 70

Tirne (minutes)

Appendix 3a. Fluorescence versus time for different substrate concentrations of methylumbelliferyl-N-acteylglucosamine (MIE-NAG). Curves are fitted with straight lines. Substrate (3 1.124 nmol) was incubated with 2 pl of standard chitobiase (Sigma Chemical Co.) in 150 pl citrate-phosphate buffer (CPB, 0.15 M, pH 5.5) until maximum fluorescence was observed. Change in fluorescence represents hydrolysis of MUF-NAG to rnethylurnbelliferone (MUF) and N-acetylglucosarnine (NAG) .

O 20 40 60 8 0 1 O0 120

Tirne (minutes)

Appendix 3 b. Fluorescence versus time for different substrate concentrations of methy lumbelli feryl-N-acte y lg lucosamine (MW-N AG) . Curves are fitted with straight lines. Substrate (48 m o l ) was incubated with 2 pl of standard chitobiase (Sigma Chemical Co.) in 150 pl citrate-phosphate buffer (CPB, 0.15 M, pH 5.5) until maximum fluorescence was observed. Change in fluorescence represents hydrolysis of MüF-NAG to methylumbelliferone (MUF) and N-acetylglucosamine (NAG).

O 20 40 60 8 O 1 O0 120 160

Time (minutes)

Appendix 3c. Fluorescence versus time for different substrate concentrations of methylurnbelliferyl-N-acteylglucosamine (MUF-NAG). Curves are fitted with straight lines. Substrate (75 nrnol) was incubated with 2 pl of standard chitobiase (Sigma Chernical Co.) in 150 pl citrate-phosphate buffer (CPB, 0.15 M, pH 5.5) until maximum fluorescence was observed. Change in fluorescence represents hydrolysis of MUF-NAG to methylumbelliferone ( N F ) and N-acetylglucosamine (N AG) .

O 2 0 4 0 60 8 0 100 120 140 160

Time (minut es)

Appendix 3d. Fluorescence versus time for different substrate concentrations of rnethylumbelliferyl-N-acteylglucosamine (MUF-NAG). Curves are fitted with straight lines. Substrate (1 00 m o l ) was incubated with 2 pl of standard chitobiase (Sigma Chernical Co.) in 150 pl citrate-phosphate buffer (CPB, 0.15 M, pH 5.5) until maximum fluorescence was observed. Change in fluorescence represents hydrolysis of MUF-NAG to methylurnbelliferone (MüF) and N-acetylglucosamine (NAG).

O 20 40 60 8 O IO0 120 140 160 180 200

Tirne (minutes)

Appendix 3e. Fluorescence versus time for different substrate concentrations of methylurnbelliferyl-N-acteylglucosamine (MUF-NAG). Curves are fitted with straight lines. Substrate (175 nmol) was incubated with 2 pl of standard chitobiase (Sigma Chernical Co.) in 150 pl citrate-phosphate buffer (CPB, 0.15 M, pH 5.5) until maximum fluorescence was observed. Change in fluorescence represents hydrolysis of MUF-NAG to methylumbelliferone (MUF) and N-acetylglucosamine (NAG).

O 50 1 O0 150 200 250

Tirne (minutes)

Appendix 3f. Fluorescence versus time for different substrate concentrations of methylumbellifery 1-N-acteylglucosamine (MUF-NAG) . Curves are fitted with straight lines. Substrate (250 nrnol) was incubated with 2 pl of standard chitobiase (Sigma Chernical Co.) in 150 pl citrate-phosphate b a e r (CPB, 0.15 M, pH 5.5) until maximum fluorescence was observed. Change in fluorescence represents hydrolysis of MUF-NAG to methylurnbelliferone (MUF) and N-acetylglucosamine (NAG).

O 5 O 1 O0 150 200 250 300

Methyhnbelliferone produced (moles)

Appendix 3g. Linear regression of fluorescence versus methylumbelliferone produced (fluorescence values detemined as maximal fluorescence fiom Appendices 3a to f)- FIuorescence values are expressed as chitobiase activity (nmol MUF liberated per unit time) as per the regression parameters: Fluorescence = -39.8 + 3.18 ( m o l e s MUF produced), r2 = 0.97.

Time (hours)

Appendix 4. Time course for decay of chitobiase (unfiltered) released into medium by individuai Daphnia magna (n=5). Bars represent standard error of mean percent chitobiase activity versus tirne. Percent chitobiase activity expressed relative to Time=O (100% activity).

Time (hours)

Appendix 5. T h e course for decay of chitobiase released into medium by individual Daphnia magna (n=6) into water that has been 0.2-pm fdtered. Bars represent standard error of mean percent chitobiase activity versus time. Percent chitobiase activity expressed relative to Time=O (1 00% activity).

O 2 4 6 8 10 12 14

T h e (days)

Appendix 6. Mean percent chitobiase activity remaining i n 0.2-prn filtered samples of incubation medium exposed to molted Daphnia magna versus time (days). Percent chitobiase activity expressed relative to Time=O (100% activity). Bars represent mean standard error (n = 12).

Body Iength (pm)

Appendix 8. Mean size fiequency distribution of subsamples (n=3) fiom Daphnia rnugna laboratory culture. Total culture volume = 10.013 L. Total subsample volume = 268 ml. Total estimated population = 1 868 individuals.

Body length (pm)

Appendix 9. Mean size fiequency distribution of subsamples (n=3), in the mixed laboratory culture of Daphnia magna - Ceriodaphnia sp. Total culture vesse1 volume = 6.934 L. Subsample volume = 268 ml. Total estimated population = 4502 individuals.

O 500 1 O00 1500 2000 2500 3000 3 500

Body Length (pm)

Appendix 10. Intermolt period versus body length for three cladoceran species. Development times @ours) for Ceriodaphnia sp. (O), Daphniaplex ( A ) , and Daphnia magna (O). (n = 20-25 for each size class). D = 26.45 + 0.02 1 (body length) ; 8 = 0.9 1, p < 0.000 1.