SEASONAL CHANGES AND TEMPERATURE INDUCED
ACTIVITY IN THE CARBOHYDRATE METABOLISM OF
THE GOLDEN SHINER, NOTEMIGONUS CRYSOLEUCAS
A Thesis
Presented to
the Faculty of the Department of Biology
East Carolina University
In Partial Fulfillment
of the Requirements for the
Degree of Master of Science in Biology
by
Nicholas A. Scandale, Jr.
May 1983
S’. X. JOVNPJK LiisHAUT
?ASX CABOLINA ÜNIVERSITT
SEASONAL CHANGES AND TEMPERATURE INDUCED
ACTIVITY IN THE CARBOHYDRATE METABOLISM OF
THE GOLDEN SHINER, NOTEMIGONUS CRYSOLEUCAS
by
Nicholas A. Scandale, Jr.
Sam N. Pennington, Ph.D.
CHAIRMAN OF THE DEPARTMENT OF BIOLOGY
DEAN OF THE GRADUATE SCHOOL
678325
ABSTRACT
Nicholas A. Scandale, Jr. SEASONAL AND TEMPERATURE INDUCED ACTIVITY IN
THE CARBOHYDRATE METABOLISM OF THE GOLDEN SHINER, NOTEMIGONUS CRYSOLEUCAS.
(Under the direction of Dr. Charles W. O'Rear, Jr.) Department of
Biology, May, 1983.
The purpose of this study was to obtain data on biochemical
activities associated with carbohydrate metabolism in the golden shiner
caused by acclimatization and expand present knowledge of the seasonal
changes in the carbohydrate metabolism.
Fish were captured using gill nets in a channelized and an unchan-
nelized stream in the Tar River water shed. Seasonal variations in
body weights, the weights of the liver, its glycogen content, and two
carbohydrate enzymes, glucose-6-phosphatase and glycogen phosphorylase,
activity values were studied. The enzymes were assayed at lO^C and
25"C to gain evidence of temperature induced activity.
In general, the liver glycogen content decreased during the study.
The phenomenon was accompanied by increased glycogen phosphorylase
activity. The fish taken from the channelized stream were often smaller
and the livers of these fish contained less glycogen. Some evidence of
temperature induced activity was found in the glucose-6-phosphatase
values. Some evidence of an inverse relationship in temperature induced
activity was found in the glycogen phosphorylase values.
ACKNOWLEDGEMENTS
I wish to give my sincere thanks and appreciation to my thesis
advisor. Dr. Charles W. O'Rear, Jr., and my thesis committee. Dr. Sam
N. Pennington, Dr. Takeru Ito, and Dr. Everett Simpson, for their advice
and criticism of this study. I would also like to thank Joey Poandl for
preparing the fine graphs and Barbara James for typing the final draft
of this study.
Laboratory facilities and equipment were provided by the Department
of Biology at East Carolina University.
DEDICATION
This study is dedicated to two very special people. My friend,
Charlotte, and my daughter, Joanie, who gave me their support and
understanding while I completed this work.
TABLE OF CONTENTS
PAGE
LIST OF TABLES . v
LIST OF APPENDIX TABLES vi
LIST OF FIGURES vii
INTRODUCTION 1
Objectives and study area selection 6
STUDY AREAS 7
Grindle Creek 7
Chi cod Creek 7
MATERIALS AND METHODS 8
Sampling 8
Glycogen content 9
Glycogen phosphorylase assay 9
Glucose-6-phosphatase assay 10
Inorganic phosphate determination 10
Water quality 11
Statistical analyses 11
RESULTS 12
Weights of the fish 12
Weight of the liver in relation to fish weight 12
Glycogen levels 12
Glucose-6-phosphatase activity 16
Glycogen phosphorylase activity 16
DISCUSSION 24
Creek differences 24
PAGE
Seasonal acclimatization 24
Temperature induced activities 26
RECOMMENDATIONS 32
LITERATURE CITED 33
APPENDIX 36
V
LIST OF TABLES
TABLE PAGE
1 Variation on the percent liver and percent glycogen
in fish taken from Chicod Creek and Grindle Creek 14
2 Variation in glucose-6-phosphatase and glycogen
phosphorylase activity in the liver of the golden
shiner (N = 5) 20
3 Variation in the percentage of active glycogen
phosphorylase in the liver of fish taken from
Chicod Creek and Grindle Creek 27
4 Comparison of water temperatures and ratios of
10“C/25‘’C assays of enzyme preparations taken from
golden shiners in Chicod Creek and Grindle Creek 30
vi
LIST OF APPENDIX TABLES
TABLE PAGE
1 Values of dissolved oxygen for Chi cod Creek and
Grindle Creek on sampling days 37
2 Values of conductivity for Chicod Creek and Grindle
Creek on sampling days 38
3 Values of pH for Chicod Creek and Grindle Creek
on sampling days 39
4 Values of dissolved nitrite for Chicod Creek and
Grindle Creek on sampling days 40
5 Values of dissolved nitrate for Chicod Creek and
Grindle Creek on sampling days 41
6 Values of dissolved ammonia for Chicod Creek
and Grindle Creek on sampling days 42
7 Values of Kjeldahl nitrogen for Chicod Creek
and Grindle Creek on sampling days 43
8 Values of ortho-phosphate for Chicod Creek
and Grindle Creek on sampling days 44
9 Values of total phosphorus for Chicod Creek
and Grindle Creek on sampling days 45
vn
LIST OF FIGURES
FIGURE PAGE
1 Metabolic map of carbohydrate metabolism
to and from glucose-6-phosphatase 4
2 Body weight of fish collected from Chicod Creek
and Grindle Creek 13
3 Variation in the percent liver in golden shiner
during spring acclimatization, based on percentage
of body weight 15
4 Variation in the glycogen content of the liver in
golden shiner, glycogen based on percent of liver mass ... 17
5 Variation in the 10®C assay of glucose-6-phosphatase
activity in the liver of golden shiner (micrograms
P04/gram liver/min.) 18
6 Variation in the ZS^C assay of glucose-6-phosphatase
activity in the liver of golden shiner (micrograms
P04/gram liver/min.) 19
7 Variation in the glycogen phosphorylase activity
(with and without c-AMP) of the liver in fish
taken from Chicod Creek 21
8 Variation in the glycogen phosphorylase activity
(with and without c-AMP) of the liver in fish
taken from Grindle Creek 22
9 Water temperature of Grindle Creek and
Chicod Creek 29
INTRODUCTION
Most research on carbohydrate metabolism has been on mammalian
species and a number of homeostatic responses to dietary or endocrine
imbalance have been reported. Studies on fish are limited to a relative
ly few species of commercial importance, such as trout, salmon, eel, cat
fish, and tuna.
Early studies of carbohydrate metabolism, in fish, were mainly con-
cerned with the compositional analyses of the tissue and eggs. Evidence
defining the process of carbohydrate metabolism was indirect and, for
the most part, inferred from various physiological and environmental
factors. The processes of glycogen metabolism are in keeping with that
worked out for mammals. Black ^ (1961) reported glycogen levels
in both muscle and liver in fish appear to be substantially lower than
those recorded for mammals. Real differences do occur and may be re-
lated to the lower body temperatures of the fish.
Little has been done to characterize the specific enzyme systems
and pathways relating to the intermediate products of carbohydrate metab
olism in fish. Macleod e^ al_. (1963) investigated glycolytic enzymes
in the tissues of the trout. Salmo gairdenerri gairdenerri, and were
the first to assay phosphofructokinase (PFK) in crude extracts of muscle
heart, brain, liver, and kidney. Evidence for the presence of all the
enzymes of the Embden-Meyerhoff glycolytic pathway except trióse phos-
phate isomerase and phosphoglycerate kinase was found. The presence of
these enzymes could be inferred from the overall glycolysis results.
Except for hexokinase and phosphofructokinase, the glycolytic enzymes
2
of the fish skeletal muscle were found to be three to one hundred times
more active than those of the liver tissues examined. Tests indicated
that PFK was the limiting enzyme (Macleod e^ aj_., 1963). Hexokinase
activity was low in all the tissues examined except heart muscle. The
results indicated not only that the fish tissues contained the same kinds
of glycolytic enzymes as are found in mammalian tissues, yeast, and
bacteria, but also that, as in glycolytic enzymes of the other species,
ADP and NAD are required. Macleod e^ al_. (1963) also determined that
the Embden-Meyerhoff pathway is not the sole pathway of degradation of
glucose in these tissues.
Williamson e;t (1967) studied glycolytic control mechanisms in
the electric organs of the electric eel, Electrophorus electricus.
Their study indicated that the principal sites of control in the glycoly-
tic pathways were glycogen phosphorylase and PFK. Control of hexose
monophosphate utilization in the electric organ was found to be similar
to that observed with mammalian systems. Their studies also demonstrated
few differences between the control sites of the glycolytic pathways of
fish and those of mammals.
There have been some recent studies of compensatory patterns of
catalytic activity during thermal acclimation. Benziger and Umminger
(1973) studied the role of hepatic glycogenolytic enzymes in salt water
adapted killifish, Fundulus heteroclitus, acclimated to ZO^C and 1.5®C.
Their investigation demonstrated that there was in increase in specific
activity of glycogen phosphorylase at the lower temperature. This in-
creased activity appeared to be responsible for the cold induced hyper-
glycemia observed. Cooper and Ferguson (1972) studied the effect of
3
cold acclimation upon glucose-6-phosphatase activity in two free living
nematodes. Their work showed that glucose-6-phosphatase activity was
greater in the cold acclimated population of Panagrellus redivivos and
relatively unchanged in Turbatrix aceti. Low and Somero (1976) studied
the adaptation of muscle pyruvate kinase to environmental temperatures
and pressures. They found most muscle pyruvate kinases exist in two
temperature dependent conformational states. The extent of conforma-
tional change which occured during catalysis was positively correlated
with all temperatures except for enzymes of deep sea species (Low and
Somero, 1976).
The seasonal differences in the carbohydrate reserves of fish,
especially those in the liver, have been known for a long time
(Plisetskaya and Kuzimina, 1972) although there are very few reports
available which concentrate on seasonal observations in nature. A
seasonal study allows a comparison to be made between recent results on
temperature acclimation and certain acclimatization phenomena (Fry,
1967; Hochachka and Somero, 1971). Recent studies (Bentley, 1976;
Carneiro and Amaral, 1983) of the hormonal control mechanism for the
carbohydrate metabolism in fish may help in handling these problems.
In this study of seasonal carbohydrate metabolism of the golden
shiner, Notemigonus crysoleucas, special attention has been paid to the
enzymes, glycogen phosphorylase and glucose-6-phosphatase, at the
glucose-6-phosphate branch point because data was available for com-
parisons with laboratory and field studies (Valtonen, 1974).
The direction of carbohydrate metabolism is determined at the
glucose-6-phosphate branch point (Fig. 1). Glycogen is directed toward
Figure 1. Metabolic map of carbohydrate metabolism to and from
glucose-6-phosphate
'i t*
GLUCOSE-1-PHOSPHATE
i’t 6 HEXOSE
GLUCOSE GLUC0SE-6-PH0SPHATE ^ MONOPHOSPHATE
I'! SHUNT
FRUCT0SE-6-PH0SPHATE
Key: 1 glycogen phosphorylase
2 glycogen synthesis
3 phosphoglucomutase
4 glucose-6-phosphatase
5 hexokinase
6 glucose-6-phosphate dehydrogenase
7 glucose phosphate isomerase
5
glucose-6-phosphate by the action of glycogen phosphorylase. At the
branch point glucose-6-phosphate can be directed toward:
(1) the Embden-Meyerhoff pathway by glucose phosphate isomerase
(2) the hexose monophosphate shunt by glucose-6-phosphate dehy-
drogenase
(3) glucose production by glucose-6-phosphatase
(4) glycogen synthesis by phosphoglucomutase and glycogen synthase.
Glucose-6-phosphatase catalyses the hydrolysis of glucose-6-phos-
phate. The enzyme is associated with the microsome fraction of cell
homogenates (Ikeda and Shimeno, 1967), although any biological advantage
of the association is not clearly apparent (Hers, 1976). Glucose-6-
phosphatase functions at an optimal pH 6.0 and is inhibited at high con-
centrations of glucose (Ikeda and Shimeno, 1967).
Glycogen phosphorylase exists in two forms, active and inactive.
Cyclic-AMP stimulates the activity of two protein kinases, one which
activates phorylase kinase, and another which converts active glycogen
synthase to inactive glycogen synthase. The activated phosphorylase
kinase converts the inactive form of glycogen phosphorylase to the
active form. This enzyme catalyzes the rate limiting step of glyco-
genolysis and indirectly controlling the activity of synthase phos-
phatase thus regulating glycogen synthesis. During glycogenolysis, the
glucose produced greatly stimulates the formation of the inactive form
of glycogen phosphorylase from the active form. The decreasing concen-
tration of the active form of glycogen phosphorylase activates glycogen
synthase which catalyzes glycogen synthesis (Hers, 1976). This antagon-
istic relationship between active glycogen phosphorylase and active
6
glycogen synthase permits a very delicate control of glycogen metabolism.
Objectives and study area selection
The objectives of this study were to obtain data on the evidence
of temperature induced activities caused by acclimatization and expand
present knowledge of the seasonal changes in the carbohydrate metabolism
of fish. The study areas, which were in the Tar River watershed, were
chosen for their proximity to each other and to Greenville.
STUDY AREAS
The study areas were Grindle Creek, which had been recently channel-
ized, and Chicod Creek, which was in a relatively natural state. Both
creeks are in the Tar River watershed.
Grindle Creek
Grindle Creek is located in the northern portion of Pitt County
originating east of Bethel and flowing in a southeasterly direction and
empties into the Tar River. Grindle Creek was almost completely chan-
nelized by a Soil Conservation watershed development project completed
in February, 1965. The project modified a total of 28.5 miles of stream
channel through snagging and clearing of 2.2 miles, enlarging 23.6 miles
of existing channel, and constructing 2.7 miles of new channel. Spoil
material was deposited on both sides of the creek.
Chicod Creek
Chicod Creek is located east of Greenville in Pitt and Beaufort
Counties. The creek flows north from its origin near Hackney to its
confluence with the Tar River about 1 mile north of Grimesland. Only
slight watershed development modifications had been made at the time of
the study. The flood plains are generally broad, wooded swamps with no
distinct channels in much of the upper reaches. Most of the Chicod
Creek is considered perennial even though there may be short periods of
no flow almost every year.
MATERIALS AND METHODS
Sampling
Sampling sites were located approximately 200 yards upstream from
the mouth of each creek. Gill nets used in the sampling included square
mesh sizes of 3 inches, 2 1/2 inches, 2 inches, 1 3/8 inches, and 1 inch.
The nets-were placed face to face in series perpendicular to stream flow
the largest mesh downstream.
The samples were taken between 0600 hours and 1200 hours on each
sampling day to eliminate variation caused by feeding. To determine
sampling days, each day of the week was assigned a number from 1 to 7
from a table of random numbers. Even numbered days were designated as
Chi cod Creek days and odd numbered days as Grindle Creek days. On the
even numbered day the first three hours of sampling time were assigned
to Chicod Creek and the last three sampling hours were assigned to
Grindle Creek and vice versa on the odd numbered days. Nets were set
January through June. The golden shiner was selected for the study be-
cause of the number of these fish captured in the creeks sampled. The
dates reported are the only dates in which golden shiners were found in
both creeks.
Date and time were recorded for all captures. The fish were frozen
in dry ice immediately after capture. They were stored in a freezer at
a temperature of -25®C. The enzyme assays and glycogen determinations
were carried out within one week of sampling. To accomplish this, the
fish were thawed just enough to allow the body cavity to be opened.
Each specimen was weighed to an accuracy of 0.01 grams and the mass of
9
the liver determined to an accuracy of one milligram.
Glycogen content
A 10% homogenate of the liver was prepared in ice cold water. A
10 microliter aliquot of the homogenate was preincubated for 10 minutes
at 55“C with 0.1 ml of 0.05 M acetate buffer, pH 4.5, 3.5 units of amylo-
glucosidase and sufficient distilled water to make a total volume of
0.5 ml. After the preincubation period, glucose was determined by the
enzymatic-colormetric glucose oxidase G0:P0 procedure using a Sigma kit.
Four milliliters of this premix was added to each sample, the glucose
standards, and reagent blank. The total volume was brought up to 4.5 ml
with distilled water. After incubation for 30 minutes at 37‘’C, the
samples were read at 500 nm on a spectrophotometer. The glucose concen-
tration for each sample was calculated from a glucose standard curve
which was determined with each set of samples. The glycogen content of
the liver was calculated as a percentage of the tissue wet weight.
Glycogen phosphorylase assay
Glycogen phosphorylase activity was measured by the release of in-
organic phosphate from glucose-l-phosphate in the presence of glycogen
and c-AMP (Hers and Hoff, 1966). Two substrates were prepared for each
assay. One substrate contained c-AMP to measure the total amount of
glycogen phosphorylase activity. The second substrate contained no
c-AMP in order to measure the amount of active glycogen phosphorylase
in the liver cells at the time of collection. They were prepared as
follows :
(1) substrate A - 0.1 M glucose-l-phosphate, 2% glycogen.
10
0.03 M c-AMP, and 0.2 M NaF
(2) substrate B - 0.1 M g1ucose-l-phosphate, 1% glycogen, and
0.2 M NaF
The pH of each substrate was adjusted to 6.1. Four tubes were prepared
for each sample. A 1% homogenate, volume of 0.05 ml, and 0.05 ml of
substrate were mixed in the bottom of centrifuge tubes and incubated at
25‘’C and 10°C for 30 minutes and 60 minutes respectively. The reaction
was stopped by the addition of 0.5 ml of 1 M trichloroacetic acid at 0
minutes (acid added before extract), 30 minutes for the 25®C assay, and
60 minutes for the 10®C assay.
G1ucose-6-phosphatase assay
G1ucose-6-phosphatase activity was based on the release of inorganic
phosphate from glucose-6-phosphate. Two tubes were prepared for each
sample. In each tube, a substrate consisting of 0.1 ml of 0.1 M glucose-
6-phosphate, pH 6.5, containing 0.002 M EDTA was mixed with 0.1 ml of a
1% liver extract. The 10°C assay was run for 60 minutes and the 25°C
assay was run for 30 minutes. The reaction was stopped by the addition
of 0.5 ml of 1 M trichloroacetic acid. The amount of inorganic phosphate
was measured at time 0 and the end of each assay.
Inorganic phosphate determination
The inorganic phosphate was determined according to the method of
Hers and Hoff (1966) by adding 0.5 ml of molybdate-sufuric acid reagent
and distilled water up to a volume of 4.8 ml. The mixture was centri-
fuged and the clear supernatent was decanted into a colormetric tube
containing a 0.2 ml of amino-naphthol sulfonic reagent. A set of
11
phosphate standards was prepared using 1.3613 g of KH2PO4 dissolved in
one liter of distilled water. A 1:10 dilution was made so that 1 ml of
standard solution corresponded to 1 micromole of phosphorus. The samples
were read at 660 nm on a spectrophotometer. The activities of the
enzymes were expressed in micrograms phosphate/gram of liver/minute.
Water quality
Water quality was monitored throughout the study for temperature,
dissolved oxygen, conductivity, pH, and nutrients. Nutrient analyses in-
eluded tests for nitrites-dissolved, nitrates-dissolved, ammonia, total
organic nitrogen-Kjeldahl, or thophosphate-dissolved, and total phos-
phorus. Samples for nutrient analyses were collected in one liter
Nalgene bottles and stored on ice until arrival at the laboratory. They
were then preserved with 1 ml mercuric chloride (40 g/1). Samples to be
analyzed for nitrites, nitrates, ammonia, and orthophosphate were fil-
tered through a 0.45 millipore filter. Analyses followed procedures
outlined in the Environmental Protection Agency's Methods for Chemical
Analysis of Water and Wastes (1971).
Statistical analyses
Data presented in the tables are in terms of means and standard
error of individual treatment groups. Statistical analyses were per-
formed using factorial analysis of variance (Sokal and Rholf, 1969).
RESULTS
Weights of the fish
The weights of the crysoleacus collected from Grindle Creek were
significantly different from those collected from Chicod Creek (P<0.01).
Generally, the fish collected from Chicod Creek weighed more than the
fish from Grindle Creek with the exception of the 4 April collection
where the difference appears minimal.
The fish in Chicod Creek showed a decline in mean weight from the
13 March collection to a minimum value on the 4 April collection rising
to a near maximum value on the 23 April. The fish from Grindle Creek
showed a rise from the minimum value on 13 March to a maximum value on
the 4 April followed by a slight decrease on 23 April. The main trends
of the fish weights are shown in figure 2.
Weight of the liver in relation to fish weight
The liver weights from Chicod Creek showed a steady decline from a
maximum on 13 March to the minimum on the 23 April. The liver weights
from Grindle Creek showed a rise from the 13 March collection to a max-
imum on the 4 April and a decline to a minimum on the 23 April (Table 1,
Fig. 3). There was a significant interaction between the collection
dates and the liver weights (P<0.05).
Glycogen levels
The glycogen levels in the fish collected from Grindle Creek were
significantly different from those collected from Chicod Creek (P<0.01).
The glycogen levels were higher in the fish from Chicod Creek, with the
Figure 2. Body weight of fish collected from Chi cod Creek and Grind!e Creek.
GBMROAADSMSYS
13 20 27 3 10 17 24
MARCH APRIL
Table 1. Variation in the percent liver and percent glycogen in fish taken from Chi cod Creek
and Grindle Creek. (N=5)
Oliver ^Glycogen
Date Collection site Mean S.E. Mean S.E.
13 March Chi cod Creek 1.770 0.0938 2.670 0.0135
Grindle Creek 0.047 0.0013 0.196 0.0121
04 April Chi cod Creek 1.230 0.0898 0.236 0.0169
Grindle Creek 1.350 0.0512 0.646 0.1940
23 April Chi cod Creek 0.472 0.0763 0.622 0.4710
Grindle Creek 0.286 0.0317 0.184 0.0273
Figure 3. Variation in the percent liver in golden shiner during spring acclimatization.
Based on percentage of body weight.
%LIVER
16
exception of the samples collected on 4 April (Fig. 4).
Glucose-6-phosphatase activity
Glucose-6-phosphatase (G6Pase) activity in the liver (Figs. 5,6,
Table 2) as measured at the assay temperatures of lO^C and 25“C is
heavily dependent on the collection dates and collection sites (P<0.005).
The 10®C assay from Grindle Creek shows a maximum value on the 13
March dropping to a minimum on the 4 April and rising again on the 23
April collection. The lO^C assay from Chi cod Creek shows little between
the 13 March and 4 April collections with a rise similar to the Grindle
Creek data on the 23 April.
The 25°C assays from Chicod Creek and Grindle Creek show little
change between the 13 March and 4 April collections with an increase to
a maximum for each creek on the 23 April collection.
All of the 25“C assay activities were higher than the 10°C assays
excluding the 13 March data for Grindle Creek.
Glycogen phosphorylase activity
The variation in the glycogen phosphorylase activity (Figs. 7,8,
Table 2) resemble that of the G6Pase only in that the 10°C and 25°C
assay temperatures are dependent on the collection dates (P<0.05).
There did not appear to be any significant differences between the creeks
on any of the collection dates. There were significant differences
between the assay temperatures (P<0.05) on each of the collection dates.
The 10°C assays from Chicod Creek and Grindle Creek show little
change between the 13 March and 4 April collection dates with a rise to
a maximum on the 23 April collection.
Figure 4. Variation in the glycogen content of the liver in golden shiner. Glycogen based on
percent of liver mass.
%GLYCOGEN
Figure 5. Variation in the 10°C assay of glucose-6-phosphatase activity in the liver of
golden shiner, (tnicrograms PO^/gram liver/min.)
MARCH APRIL
Figure 6. Variation in the 25°C assay of glucose-6-phosphatase activity in the liver of
golden shiner, (micrograms PO^/gram liver/min.)
GA-C6T-PIVAISTEY /f(mPl^icvmaro0gemrian4mr.s)
MARCH APRIL
Table 2. Variation in glucose-6-phosphatase and glycogen phosphorylase activity in the liver
of the golden shiner. (N=5)
Glucose-6-phosphatase Glycogen Phosphoryl ase
10°C 25°C 10°C 25°C
Date Collection Site Mean SE Mean SE Mean SE Mean SE
13 March Chicod Creek 5.82 0.09 36.29 3.24 43.75 10.95 28.28 4.09
Grindle Creek 112.63 0.90 5.31 0.21 46.72 3.49 83.13 5.10
04 April Chicod Creek 0.26 0.03 8.70 0.03 53.84 7.15 35.90 2.15
Grindle Creek 0.23 0.08 9.36 2.75 31.49 4.93 2.80 1.41
23 April Chicod Creek 74.26 20.70 109.43 17.19 195.92 66.00 40.87 19.22
Grindle Creek 111.31 7.14 207.35 28.21 266.29 77.70 134.83 28.30
Figure 7. Variation in the glycogen phosphorylase activity (with and without c-AMP) of the liver
in fish taken from Chicod Creek.
25*C ASSAY
AGSPHCLOYSTCPHIEOVORGIYTELAYN (mlPivic0er4or/g/mrraamiins.)
MARCH APRIL MARCH roAPRIL
Figure 8. Variation in the glycogen phosphorylase activity (with and without c-AMP) of the liver in
fish taken from Grindle Creek.
10® C ASSAY 25® C ASSAY
AGPHCLOYSTPCHIVORGYITLEASYNE (Plmiv0ic4er/orgg/rmraomins.)
13 20 27 3 10 17 24
march APRIL MARCH APRIL
23
All of the 10°C assays from both creeks were higher than those of
the 25®C assays with the exception of the 25®C assay from Grindle Creek
on the 13 March.
DISCUSSION
Creek differences
The weights of the fish taken from Grindle Creek indicate that
these fish are smaller than those taken from Chicod Creek. This con-
si stent with reports by Bayless and Smith (1964) who reported a 90%
reduction in weights of fish taken from channelized streams.
The greatest single factor affecting a fish population appears to
be the amount of stream cover (Tarplee e^ , 1971). The stream canopy
is an important source of organic input into aquatic systems (Hall,
1972). Loss of in-stream shelter and feeding areas may contribute to
changes in size characteristics and weights of fish within a population.
The loss of in-stream shelter and feeding areas implies a poor nutritional
status for fish taken from channelized streams. The low levels of
glycogen observed in fish taken from Grindle Creek are similar to gly-
cogen levels in catfish, Ictaluras punctatus, after fasting for thirty
days. The glycogen levels of fish taken from Chicod Creek compare
favorably to the glycogen levels in catfish which have been feeding
(Ottolenghi e;t ^., 1981). It is known that the glycogen content of
the liver is influenced by nutritional status (Black et ^., 1961;
Hochachka, 1961). The low levels of glycogen, observed in Grindle
Creek, are consistent with these observations.
Seasonal acclimatization
It has been known for some time that the glycogen content of the
liver in fish is maintained at a higher level during acclimatization
to cold waters in winter than warm waters in summer (Plisetskaya and
25
Kuzmina, 1972). DeVlaming and Pardo (1975) reported increased glycogen
deposition in liver slices, taken from the golden shiner, maintained at
12®C. High glycogen levels occur in the livers of fish found in Chicod
Creek during March. Spawning occured during the April collections be-
cause females ripe with eggs were collected in both creeks. The rapid
decline in glycogen levels in the fish taken from Chicod Creek agree
with the reports by Bullock (1954) and Black (1961) concerning
changes in the liver glycogen during spawning. The glycogen decrease
is closely connected with the production of eggs and sperm (Bullock,
1954). The fish collected from Grindle Creek did not show the dramatic
decline in glycogen levels. In fact, there was a slight increase in
glycogen levels during the month of April. This may reflect an increase
in food supply in the creek (Mayerles and Butler, 1971), although there
is no evidence to support this. It is possible that the fish collected
during the spawning period may have moved into Grindle Creek from the
Tar River. The phenomena in question are closely connected with accli-
matization, though Hochachka and Somero (1971) do not accept high gly-
cogen synthesis as an obligatory feature in acclimatization to cold.
When the enzyme activities studied are considered against the back-
ground described above, the changes are most interesting. Glucose-6-
phosphatase activity declined during spawning in early April and in-
creased slightly late in April. This enzyme, which liberates glucose in-
to the blood both by gluconeogenesis and glycogenolysis resembles the
corresponding enzyme in mammalian liver and is controlled through meta-
bolites (Hers, 1976), glucose (Ikeda and Shimeno, 1967), and hormonally
(DeVlaming and Pardo, 1975). The low activity, while there was a
26
plentiful supply of food in the stomach during April may be understood
by reference to the control of insulin on absorbed glucose in the blood.
Insulin regulates the carbohydrate metabolism of the teleost by hor-
monal mechanisms similar to those operating in mammals (Carneiro and
Amaral, 1983). Golden shiners adapted to show a net decline in
liver glycogen even though insulin promoted glycogen deposition at the
same temperature (DeVlaming and Pardo, 1975). This observation may ex-
plain the slight increase in glycogen phosphorylase activity during the
same period.
On 13 March, 73% of the available glycogen phosphorylase was active
in the fish taken from Chicod Creek. This increased to 90% on 4 April
and decreased to 50% by 23 April. This increased activity is responsible
for the rapidly declining glycogen content in the livers of these fish
(Fig. 4). Only 35% of the available glycogen phosphorylase was active
in fish taken from Grindle Creek on 13 March (Table 3). The low levels
of glycogen found in these fish indicate that the energy reserve of
these fish is nearly depleted. The decrease in the available glycogen
phosphorylase in the livers of fish collected on 4 April may be respon-
sible for the increased glycogen levels in the livers of these fish.
As spawning activity increased, the total amount of glycogen phosphory-
lase in fish from Grindle Creek increased. 89% of this enzyme was active
causing a decline in the glycogen content in the livers of the fish
collected from this creek on 23 April.
Temperature induced activities
The data presented favors the view that temperature induced
rabie 3. Variation in the mean percentage of active glycogen phosphorylase in the liver of fish
taken from Chi cod Creek and Grindle Creek.
Date Collection Site 10°C Assay 25°C Assay
13 March Chi cod Creek 0.74 0.52
Grindle Creek 0.35 0.44
04 April Chicod Creek 0.90 0.69
Grindle Creek 0.82 0.11
23 April Chicod Creek 0.50 0.28
Grindle Creek 0.89 0.58
28
compensation of enzyme activity takes place in the liver of the golden
shiner, Notemigonus crysoleucas. The ratios of enzyme activities incu-
bated at 10®C: incubated at 25°C seems to be connected with regulation
of isozyme systems (Hochachka and Somero, 1971). Hochachka and Somero
suggested three mechanisms which might be responsible for regulating
the rates of enzyme activity after period of acclimation. Organisms
acclimate to temperatures by:
(1) increasing the concentrations of certain key enzymes
(2) generating new isozymes ideally suited to function in the
new thermal environment, or
(3) altering the microenvironment within which the enzyme
functions, thus producing compensated rates of activity by
modulation of enzyme efficiency rather than quantity.
The most striking evidence for this temperature induced activity
is the data from 13 March. The temperature difference between the
creeks was greatest on that day (Fig. 9). The high ratios of the 10®C
assays: 25°C assays of qlucose-6-phosphatase activity in the livers of
fish from Grindle Creek contrast sharply with the low ratios observed
from Chicod Creek (Table 3). The low ratios observed in April, when
the water was warm in both creeks, adds support to this observation.
Cooper and Ferguson (1972) reported the same kind of activity for
glucose-6-phosphatase in the nematode, Panagrellus redivivos.
The low ratios of glycogen phosphorylase activities incubated at
10°C: incubated at 25'’C in March followed by high ratios during April
demonstrate temperature modulated activity (Table 4). In this case,
there is an inverse temperature affect. This observation is supported
Figure 9. Water temperatures of Grindle Creek and Chicod Creek.
Table 4 . Comparison of water temperatures and ratios of 10°C/25°C assays of enzyme preparations
taken from golden shiners in Chicod Creek and Grindle Creek.
G1ucose-6-phosphatase Glycogen Phosphorylase
Date Collection site Water Temp.(°C) 10°C/25Oc assay 10°C/25°C assay
13 March Chicod Creek 16.0 0.16 1.55
Grindle Creek 9.0 21.21 0.56
04 April Chicod Creek 18.0 0.03 1.50
Grindle Creek 19.5 0.02 11.25
23 April Chicod Creek 18.5 0.68 4.79
Grindle Creek 19.0 0.54 1.98
CO
o
31
by reports from Hazel and Prosser (1974) and Hochachka and Somero
(1971).
RECOMMENDATIONS
The small sample sizes and limited time of this study suggest that
it be used as a preliminary report for the further study of the ecology
and biochemistry of the teleost, Notemigonus crysoleucas.
Several questions have brought to mind as a result of this study.
First, the question of seasonal carbohydrate metabolism. This report
cataloged information dealing with only a small phase in the life cycle
of the golden shiner, that of spawning and acclimatization from winter
to spring. A year long study would give a much clearer picture of the
ecological aspects of the life cycle of this fish population.
Secondly, this report indicates the presence of temperature in-
ducible systems for both glucose-6-phosphatase and glycogen phosphorylase.
Studies need to be carried out to determine the characteristics of each
of the isozyme systems.
It would be advantageous to study the other metabolic pathways
leading from the glucose-6-phosphate branch point. This would give a
more complete picture of carbohydrate metabolism.
Additional studies concerning the control mechanisms for glycogen
deposition and degradation should be carried out. These would shed
more light on the importance of glycogen as a source of energy to the
golden shiner.
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APPENDIX
37
Appendix Table 1. Values of dissolved oxygen for Chicod Creek and Grindle
Creek nn samnlino davs. ÍMG/L)
Date Chicod Creek Grindle creek
13 March 9.25 10.18
04 April 8.32 10.65
23 April 9.50 10.73
38
Appendix Table 2. Values of conductivity for Chicod Creek and Grindle
Creek on samolinq days. (yMHOS/CM^)
Date Chicod Creek Grindle Creek
13 March 45.16 62.53
04 April 87.50 92.52
23 April 112.43 145.34
39
Appendix Table 3. Values of pH for Chi cod creek and Grindle creek on
sampling days.
Date Chi cod Creek Grindle Creek
13 March 5.0 5.7
04 April 6.5 5.3
23 April 5.6 5.9
Appendix Table 4 . Values of dissolved nitrite for Chicod creek and
Grindle Creek on sampling days. (MG/L)
Date Chicod Creek Grindle Creek
13 March 0.010 0.009
04 April 0.014 0.011
23 April 0.009' 0.013
Appendix Table 5 Values of dissolved nitrate for Chi cod creek and
Grindle creek on sampling days. (MG/Lj
Date Chi cod Creek Grindle Creek
13 March 0.408 0.467
04 April 0.300 2.100
23 April 0.500 2.400
42
Appendix Table 6. Values of dissolved ammonia for Chicod Creek and
Grindle Creek on sampling days.(MG/L)
Date Chicod Creek Grindle Creek
13 March 0.017 0.058
04 April 0.233 0.033
23 April 0.795 0.495
Appendix Table 7. Values of Kjeldahl nitrogen for Chicod Creek and
Grindle creek on sampling days. (MG/l)
Date Chicod Creek Grindle Creek
13 March 1.44 1.46
04 April 2.80 2.00
23 April 4.08 4.71
44
Appendix Table 8. Values of ortho-phosphate for Chicod Creek and Grindle
Creek on sampling days. (MG/L)
Date Chicod Creek Grindle Creek
13 March 0.006 0.006
04 April 0.048 0.025
23 April 0.146 0.250
Appendix Table 9. Values of total phosphorous for Chicod Creek and
Grindle creek on sampling days. (MG/L)
Date Chicod Creek Grindle Creek
13 March 0.138 0.154
04 April 0.183 0.115
23 April 0.215 0.319