Biochemical Responses to Hypoxia in
the Long-Jawed Mudsucker I:
Metabolites
Scott Stonington
Hopkins Marine Station
Issue in Marine Biology: 175H
June 1999
Advisor: George Somero
Permission is granted to Stanford University to use the
abstract and citation of this paper
Abstract
Gillichthys mirabilis (the long-jawed mudsucker, Family Gobiidae), acclimated to 14°C, were exposed to
different periods of hypoxia. I selected the hypoxia intensity based on critical oxygen concentration (Pai)
data obtained from three fish, all of whom displayed Pait of 1.0 mg/L. I placed experimental fish in a
consistent oxygen concentration of 0.8 mg/L, and sacrificed them after 0, 8, 24 and 72 hours of exposure by
freeze-clamping in liquid nitrogen. I examined glycogen, lactate and ethanol concentrations in white
muscle, liver, heart and brain. I found no ethanol in any tissue. Lactate accumulated significantly and
similarly in all tissues throughout the time course. Glycogen changed significantly only in liver, which had
very large initial stores (6.87 + 1.46 mg/g), and drastically depleted stores by 8 hours (0.59 + 0.22 mg/g)
White muscle had consistently low glycogen (mean 0.27 + 0.01 mg/g). Brain showed some hints of
glycogen depletion (p = 0.053) by 72 hours. I conclude that during hypoxia in Gillichthys mirabilis,
glycogen is mobilized from stores in the liver and transported to meet the energy demands of brain, heart
and white muscle, which conserve their own glycogen.
Introduction
Gillichthys mirabilis is a teleost fish of the family Gobiidae that has successfully adapted
to a wide range of extreme environments. Possibly the most prominent of these extremes are the
hypoxic and sulphurous sloughs of Western California - plains of partially toxic mud with tidal
drainages that leave the substrate exposed to air and sun. To escape this exposure, Gillichthys
mirabilis have been shown to hide in small burrows created by invertebrates in the slough's
muddy substrate (Congleton, 1974). They thus spend significant time in small volumes of water
that may experience extreme hypoxia and fluctuating temperatures. Indeed, as long ago as 1930,
researchers became interested in Gillichthys mirabilis's ability to acclimate to temperature
fluctuations (Sumner and Doudoroff, 1938), and more recently the gobiid has been used as a
model for both heat and osmotic shock tolerance (Lin and Somero, 1995; Kultz and Somero,
1995; Kultz 1995; Kultz and Somero, 1994). The Gillichthys mirabilis habitat, however, is also a
site of periodic extreme hypoxia, and they may thus prove valuable as a model for studying
hypoxia tolerance.
Hypoxia is an environmental stress pertinent to almost all aerobic organisms. It generally
falls under two categories: environmental and physiological hypoxia. Environmental hypoxia
occurs when an organism cannot obtain enough oxygen from its environment to maintain normal
systemic energy consumption. It thus strikes at a systemic level, and the organism must address
the problem in an integrated fashion among all tissues. In physiological hypoxia, despite
abundant environmental oxygen, a tissue's energy demand is greater than the rate of oxygen
supply to that tissue. This is usually a result of high activity, where one tissue (usually muscle)
must work at such a rate that it cannot produce ATP quickly enough through aerobic pathways
These two forms of hypoxia present very different problems to an organism. Moreover,
hypoxia can span a large range of severity, from anoxia to common fluctuations in oxygen
transport throughout the body. It is thus understandable that organisms might develop different
strategies to different severities and forms of hypoxia. These strategies generally fall under three
categories. First, organisms may reduce their physiological and behavioral activity to decrease
the demand for ATP, and thus for oxygen (Hochachka et al., 1996). This usually involves a
suppression of non-essential functions. Second, organisms may attempt to increase extraction of
oxygen from the environment and delivery of oxygen to hypoxic tissues. This can include
hyperventilation, vascularization of hypoxic tissues or alteration of blood properties such as
hematocrit or hemoglobin concentrations. Third, and possibly most prominently, organisms may
revert to anaerobic metabolic pathways to supply their remaining ATP needs. Consequently
since anaerobic metabolism is far less efficient than aerobic metabolism, organisms require a
greater quantity of metabolic substrate per ATP output, and thus may need to mobilize energy
stores such as glycogen. Moreover, anaerobic metabolism produces undesirable end-products,
such as lactic acid, which can cause acidotic damage to tissues. Thus tissues with high metabolic
costs may be in greater danger from hypoxia than tissues that operate at lower metabolic output.
To protect against acidotic buildup and to mobilize energy stores, organisms may employ tissue¬
specific metabolic strategies, shuttling energy-substrates throughout the body to provide for
various tissues' energy needs.
The purpose of this investigation is to discern which of these strategies Gillichthys
mirabilis employs to resist the periods of environmental hypoxia in its environment. A logical
first place to turn, of course, is to the regulation of metabolites, since any change in metabolic
strategy should alter both the use of energy stores and the quantity and form of waste products.
Metabolic substrates and end-products can be used as markers for the "black box" of overall
metabolic strategy. The most obvious of these markers are glycogen and lactate, because
glycogen is the primary energy-storage molecule in vertebrates and the lactic-acid-producing
metabolic pathway, glycolysis, is the most common of anaerobic strategies. There are of course,
numerous exceptions to this. Goldfish, for instance, which experience very long periods of
hypoxia, employ a metabolic pathway that produces ethanol, which is then excreted as waste
(Johnston and Bernard, 1982). Organisms may also deplete energy stores other than glycogen,
such as fatty acids or proteins. Alternatively, they may maintain a high constitutive anaerobic
throughput, and thus display little change in anaerobic metabolism during hypoxia. Nonetheless
glycogen and lactate are invaluable markers that can help elucidate the overall metabolic strategy
that Gillichthys mirabilis use to survive their hypoxic slough.
Materials and Methods
I conducted all experiments in collaboration with a colleague, Joshua Troll, who used the
same test population and tissues for assays on glycolytic enzyme activity. Gillichthys mirabilis
were collected from a lagoon at the University of California Santa Barbara and acclimated to
14°C for several months. On three separate occasions after their acclimation, we placed
individual fish in a sealed tank coupled to an oxygen electrode to measure the fish's rate of
oxygen uptake versus available oxygen concentration. These data are shown in figure 1 and were
used to determine the Pert of our stock of fish (the oxygen concentration below which an
organism's oxygen consumption rate begins to decline). We found the Pent to be approximately
1.0 mg/L, and designed the remainder of our experiments appropriately.
We placed four sets of seven fish (3.5g-19.25g) in aerated Tupper-Ware"" containers in a
tank liked to a nitrogen-gas hypoxia setup. Containers were used to facilitate sacrificing the fish
without causing any substantial handling stress which might alter the fish's metabolic state. Äfter
one day in this new setting, we sacrificed one of the four sets of fish and reduced the oxygen
concentration in the tank to 0.8 mg/L, 80% of their Perit. The remaining three sets of fish were
sacrificed after 8, 24 and 72 hours of hypoxia. Data for this exposure are shown in figure 1.
We sacraficed the fish by freeze-clamping them in liquid nitrogen to immediately
preserve their tissues and prevent both post-mortem enzyme activity and metabolite breakdown.
We transferred the frozen fish to storage at -80°C until removing them for dissection. During the
dissections, we kept the fish on dry ice, allowing us to perform separate dissections and return the
fish to storage without disturbing their endogenous enzyme integrity by freezing and thawing
their tissue. This technique allowed us to remove white muscle at one sitting (-400mg), and liver
(100mg), heart (-40mg) and brain (-20mg) at the next. We parceled these tissues into two sets,
one for endpoint-assays (this paper), another for enzyme activity assays (see data from Joshua
Troll, this volume). I placed tissues for endpoint assays immediately in 0.6 N perchloric acid (5
ul/mg tissue) to arrest endogenous enzyme activity. I then homogenized these mixtures with an
Ultraturax?.
For the four tissues mentioned above, I assayed for glycogen according to the method of
Keppler and Decker (1969). Two aliquots of homogenate from each tissue of each fish were
neutralized to protect assay enzymes from deproteinization. To one of these, I added 250 ul of
11.0 units/ml amyloglucosidase solution in 0.2 M acetate buffer (pH 4.8) to hydrolyze the tissue's
glycogen into glucose. The other aliquot (with 250 ul acetate) served as a blank of free glucose
against which glycogen data could be compared. After digestion, I centrifuged the samples and
used three 20 ul aliquots of supernatant for triplicate assays. I added these samples to 100 ul of
reaction mixture (1 mM ATP; 0.9 mM B-NADP ; 5 ug glucose-6-phosphate dehydrogenase/ml.),
and measured their change in absorbance before and after reaction with hexokinase. Absorbances
were measured at 340 nm on a SpectraMax 340c 96-well spectrophotometer using SOFTMAX¬
PRO software, all manufactured by Molecular Devices. The absorbances from triplicate wells
were averaged to obtain a value for a single tissue, and actual glucose concentration values were
derived using the numerical analysis detailed in Keppler and Decker (1975)
I assayed for l-lactic acid and ethanol using the methods and industrial kits provided by
Boehringer Mannheim (Cat. Nos. 139084 and 176290). For each assay, I added three 20 ul
aliquots of neutralized homogenate solution to wells containing 100 ul reaction solution
(provided by the Boehringer Mannheim kits). To develop a standard curve for absorption change
versus lactate or ethanol concentration, I also added aliquots of lactate and ethanol standard
solution (of 9.7 mg/L, 29.1 mg/L, and 48.5 mg/L for lactate; 4.5 mg/L, 13.5 mg/L, 22.5 mg/L for
ethanol). I then measured the absorbance of these solutions before and after reaction with lactate
dehydrogenase and alcohol dehydrogenase, respectively. The method for these measurements is
described above in the glycogen assay.
-Results
Oxygen consumption rate data as a function of oxygen concentration for three fish are
shown in figure 1. This graph provides information both on the amount of oxygen that
Gillichthys mirabilis consume as well as the critical oxygen concentration below which they
begin to lose their ability to draw oxygen from their surroundings (Perit). Pent is thus measured as
the oxygen concentration below which oxygen consumption rate consistently declines. The Perit
for all three experimental fish was near 1.0 mg/L, and was independent of body mass.
No ethanol was discernable in any of the four tissues. Ethanol is a volatile compound,
and if present in small quantities in the tissues, may not survive in substantial quantities
throughout the homogenizations and dilutions in the assay. The data, however, are clearly
negative and rule out the existence of large ethanol concentrations in any of the tissues.
Figures 3-6 show the effects of hypoxia on both glycogen and lactate concentration in
each tissue. In all four tissues, lactate rose significantly after 8 hours of hypoxia (figure 7)
(pS.001), and in liver and heart it rose again by 72 hours (pS.001). Each tissue’s increase in
lactate is generally correlated with initial lactate levels, which were consistently highest in the
brain and lowest in the liver (pS.005).
Significant changes in glycogen concentration appeared only in liver (pS.001), which had
a comparatively large initial store of glycogen that was almost completely depleted by 8 hours.
The liver’s mean initial glycogen store (6.87 mg/g) was four times the store in the next highest
tissue, heart (1.68 mg/g) and twenty-four times that in white muscle (0.29 mg/g). This low white
muscle glycogen was consistent throughout the time-course. The observed decrease in brain
glycogen was nearly significant (pS.053) at seventy-two hours, hinting at a pattern, but not
significant enough to draw any conclusions.
Discussion
The 1.0 mg O/L Pait that I observed in G. mirabilis is not surprising given gobiid fishes
capacity for hypoxia tolerance. Critical oxygen concentrations of 0.6 mg/L, for example, have
been observed in gobies of the genus Typhlogobius. Interestingly, however, the Pent values
observed in my study are on the low end of previously measured Pert in G. mirabilis, which range
from 1.0-1.5 mg/L (Congleton, 1974). This may indicate that the G. mirabilis from my
experimental population are more tolerant of hypoxia than most populations of the species. In a
broader sense, the Pent values are informative in comparison to values from other organisms and
to the oxygen concentrations found in most marine habitats. The oxygen concentration in surface
seawater ranges from 8.1 mg/L to 9.5 mg/L (Yang et al., 1992). The Pet of yellowfin tuna, a
highly aerobic pelagic fish, is 4-5 mg/L (Barbara Block, personal correspondence). Even
Scorpaena guttata, a benthic fish that lives at 180m where oxygen concentration is one third of
surface sea water, has a Pent of 3.0 mg/L (Yang et al., 1992). One of S. guttata's confamilials
(Sebastolobus alascanus), however, lives in the oxygen minimum zone and has a Pert equivalent
to that of G. mirabilis (Yang et al., 1992). Thus G. mirabilis has developed a physiology which
facilitates life under hypoxic stress of similar intensity to that of the oxygen minimum zone.
The glycogen levels that I found in G. mirabilis, especially those in the liver, shed
important light on the G. mirabilis strategy for hypoxia tolerance. Glycogen levels changed
significantly only in the liver, where large initial stores underwent a twelve-fold depletion by
eight hours of hypoxia. Lactate, however, accumulated in all four organs, indicating that
glycolysis proceeded according to each tissue's energy demand. Moreover, lactate levels were
the lowest in liver, indicating that the glucose formed by breakdown of glycogen was not being
consumed there, but was transported from the liver to other tissues. Lactate levels were highest in
the brain and heart, neither of which depleted their own glycogen, indicating that they were
receiving energy input from another source. These data argue that, in response to hypoxia, G.
mirabilis may mobilize energy stores in the liver and shuttle glucose to support the energy
demands of other tissues. These fish may thus be integrating each tissue's energy resources and
demands in response to their systemic hypoxic stress.
This strategy contrasts with anaerobic systems in other organisms. Although goldfish, for
example, deplete their liver glycogen and maintain white muscle stores in response to anoxia
(Thillart et al., 1980), they also deplete between 75 and 95% of their brain glycogen within two
hours of exposure to anoxia (Schmidt and Wegner, 1988). This strategy is primarily in response
to anoxia, however, and it is possible that G. mirabilis conserve their brain glycogen until the last
moment or mobilize it under more severe hypoxia. Moreover, G. mirabilis does not tolerate
anoxia to the same degree as goldfish, and anoxia-tolerance may entail a mechanism distinct from
hypoxia-tolerance and thus be irrelevant in comparison to G. mirabilis.
Another interesting contrast to other organisms' physiology is the size of G. mirabilis's
glycogen stores. The initial stores of glycogen in the liver (6.87 + 1.46 mg/g) are larger than in
most vertebrates. Human livers typically have 0.1-1.0 mg/g glycogen (Johnson and Fusaro, 1966;
Lee and Whelan, 1966). Kepler and Decker (1969), who developed the glycogen assay employed
in this study, found rat liver glycogen at 0.55 +0.1 mg/g. Glycogen levels in some other
hypoxia-tolerant vertebrates, however, are much larger than those observed in this study.
Glycogen makes up 30% of the crucian carp liver (300 mg/g), and the freshwater turtle has liver
glycogen 25 times the quantities observed here (Lutz and Nilsson, 1997). Moreover, although the
glycogen levels found here in the G. mirabilis brain (1.15 +0.11 mg/g) are 3-fold larger than in
rat, mouse and rainbow trout, they are nonetheless 2-3-fold smaller than in the crucian carp,
goldfish, and freshwater turtle (Lutz and Nilsson, 1994). This difference between G. mirabilis and
the carp and turtle may be due to the types of hypoxia that each organism encounters in its natural
environment. Carp and freshwater turtles are accustomed to longer durations of hypoxia (weeks
to months), and may thus need to mobilize greater glycogen stores between opportunities for
gluconeogenesis. Gillichthys mirabilis, on the other hand, have the tides to usher in fresh volumes
of oxygenated water every twelve hours or less, so smaller glycogen stores may be sufficient to
sustain them during hypoxic period at low tide.
Different time-courses of hypoxia may also explain the failure to detect ethanol in G.
mirabilis tissues. An ethanol-producing metabolic pathway appears in carp and goldfish to
prevent acidotic buildup from lactic acied during very long periods of hypoxia (Johnston and
Bernard, 1983). The absence of ethanol-producing pathways in G. mirabilis may indicate, in
part, that acid accumulation is less of a problem over twelve-hour periods than over longer
exposures. Gillichthys mirabilis, both in metabolic pathway strategies and in glycogen
concentrations, appear to lie on a gradient dictated by the severity of hypoxia found in the natural
environment.
Certain features of the G. mirabilis biochemistry, however, are in contrast to this apparent
trend. Glycogen in white muscle, for instance, was remarkably lower than in most vertebrates,
including both anoxia-tolerant and anoxia-sensitive species. It remained at a constant mean of O.3
mg/g, an order of magnitude smaller than in goldfish (Thillart et al., 1980) and two orders of
magnitude smaller than in humans (Hultman, 1967). Such low white muscle glycogen may be
due to the limited capacity of G. mirabilis for intense "burst" locomotion (personal observation).
A second contrast to other hypoxia-tolerant species is the consistent accumulation of lactate in G.
mirabilis. Normoxia-acclimated goldfish show no significant lactate accumulation under hypoxia
in white muscle or red muscle, but show three-fold increases in liver (Thillart et al., 1980). In G.
mirabilis, on the other hand, lactate accumulated in all tissues, with brain lactate consistently
highest and liver lactate consistently lowest. High lactate concentrations in the brain indicate a
high mass-specific metabolic rate, which is typical of vertebrate brain tissue, but the low lactate
concentrations in the liver are surprising because vertebrate livers normally have high mass-
specific metabolic rates (Hulbert and Else, 1981). This difference may indicate that the liver
enters metabolic arrest during hypoxia, suppressing certain functions that normally consume so
much energy. Indeed, goldfish have been shown to arrest protein synthesis in the liver during
anoxia (Smith et al., 1996). Alternatively, the low lactate levels in liver could be due to the
liver's role in removing lactate from the blood stream. The liver could produce lactate and
subsequently process it, thereby containing low lactate levels but nonetheless conducting
glycolysis and gluconeogenesis. Such processing, however, requires energy that may not be
available during hypoxia, and it would seem counter-productive to burn glycogen and
subsequently spend energy rebuilding storage substrates while still under hypoxic stress.
This study, therefore, illustrates at least two strategies that G. mirabilis may employ in
response to hypoxic stress. First, I conclude that under hypoxic conditions, G. mirabilis rely on
glycolysis for ATP production, and supply the necessary substrates for glycolytic flux by
depleting liver glycogen stores and shuttling them as glucose to other tissues. Second, G.
mirabilis may metabolically suppress the liver so that it consumes less ATP and produces less
lactate than other tissues. These strategies are a clear example of a complex systemic response to
environmental hypoxia, one that integrates the needs and responses of different tissues into a
unified strategy for the whole organism's survival.
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(Teleostei: Gobiidae) and some related gobies. Biol. Bull. 146: 186-205.
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Johnson, J.A. & Fusaro, R.M. (1966). The quantitative enzymic determination of animal liver
glycogen. Anal. Biochem. 15: 140-149. 1966;
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Lutz, P.L. & Nilsson, G.E. (1994). The Brain without Oxygen, Causes of Failure and
Mechanisms for Survival. Austin, TX: R.G. Landes.
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Schmidt, H. & Wegner, G. (1988). Glycogen phosphorylase in fish brain (Carassius auratus)
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Figure 1. Critical oxygen concentration (Pqu) for three Gillichthys mirabilis. Pait is taken as the
oxygen concentration below which the fish begin to lose their ability to draw oxygen
from their surroundings (here an upward slope). All three fish have a Peri of 1.0 mg/L.
This value was used to determine the experimental oxygen concentration (0.8 mg/L).
Figure 2. Lactate and glycogen concentrations in white muscle of fish exposed to 0, 8, 24, and
72 hours of 0.8 mg O/L hypoxia. Error bars represent onestandard deviation from the
mean of each time-point. (*) mark significant changes from control (O hours).
Significance was determined using a standard one-way ANOVA (p.001) with a
Tukey's test (HSD = 0.54). Sample sizes are given in parenthesis on the x-axis.
Figure 3. Lactate and glycogen concentrations in heart of fish exposed to 0, 8, 24, and 72 hours
of 0.8 mg O/L hypoxia. Error bars represent one standard deviation from the mean.
*) mark significant changes from control (0 hours). (**) mark significant changes
from 8 hours. Significance was determined using a standard one-way ANOVA
(pS.001) and a Tukey's test (HSD = 0.58). Sample sizes are given in parentheses on the
X-axis.
Figure 4. Lactate and glycogen concentrations in brain of fish exposed to 0, 8, 24, and 72 hours
of 0.8 mg O/L hypoxia. Error bars represent one standard deviation from the mean.
(*) mark significant changes from control (0 hours). (**) mark significant changes
from 8 hours. Significance was determined using a standard one-way ANOVA
(pS.001) and a Tukey's test (HSD = 0.52). Sample sizes are given in parentheses on the
X-axis.
Figure 5. Lactate and glycogen concentrations in liver of fish exposed to 0, 8, 24, and 72 hours
of 0.8 mg O/L hypoxia. Error bars represent one standard deviation from the mean
(*) mark significant changes from control (0 hours). (**) mark significant changes
from 8 hours. Significance was determined using a standard one-way ANOVA (p..001
for lactate; p5.0001 for glycogen) and a Tukey's test (HSD - 0.31 for lactate; HSD-
2.87 for glycogen). Sample sizes are given in parentheses on the x-axis.
Figure 6. Glycogen concentrations in liver, heart, white muscle, and brain of fish exposed to 0,
8, 24, and 72 hours of 0.8 mg O/L hypoxia. Error bars represent one standard
deviation from the mean. Glycogen concentration changes significantly only in liver.
Figure 7. Lactate concentrations in liver, heart, white muscle and brain of fish exposed to 0, 8,
24, and 72 hours of 0.8 mgO/L hypoxia. Error bars represent one standard deviation
from the mean. Lactate accumulates significantly in all tissues throughout the time¬
course.
100
40
20
Figure 1.
—e— Fish 1 (4.92g)
—0— Fish 2 (17.479)
—v— Fish 3 (25.089)


.


Vtet

00 0.5 1.0 1.5 2.0 2.5 3.0 3.5 4.0 4.5 5.0 5.5 6.0 6.5 7.0
O, Concentration (mg/L)
1.8 -
1.5
0.9
0.6
0.3
0.0 +

Glycogen
— Lactate
O Hrs (7)
Figure 2.
8 Hrs (7) 24 Hrs (6) 72 Hrs (6)
Hypoxia Duration
4.0
3.5
3.0 -
- 25
2.0
1.5
1.0
0.5
0.0-
Glycogen
□ Lactate
O Hrs (7)
Figure 3.
8 Hrs (7) 24 Hrs (6)
Hypoxia Duration
72 Hrs
2.0
1.0
0.5
0.0 -
— Glycogen
— Lactate
O Hrs (7)
Figure 4.
24 Hrs (6)
8 Hrs (7)
Hypoxia Duration
72 Hrs (6)
O Hrs (7)
Figure 5.
Glycoger
Lactate

8 Hrs (7) 24 Hrs (6) 72 Hrs (6)
Hypoxia Duration
10
0 -

O Hrs (7)
Figure 6.
I I
8 Hrs (7) 24 Hrs (6)
Hypoxia Duration
Liver
— Brain
Heart
□ White Muscle
ME
72 Hrs (6)
0 +
Liver
White Muscle
Heart
— Brain

O Hrs (7)
Figure 7.

8 Hrs (7) 24 Hrs (6)
Hypoxia Duration
72 Hrs (6)