PHYSIOLOGICAL EFFECTS OF WATER STRESS ON
TWO SPECIES OF HETEROMYID RODENTS
by
JACQUELINE A. HOMAN, B.S., M.S.
A DISSERTATION
IN
ZOOLOGY
Submitted to the Graduate Faculty of
Texas Tech University
in Partial Fulfillment of
the Requirements for
the Degree of
DOCTOR OF PHILOSOPHY
Approved
May, 1992
/ /2 el'? 3
Copyright © 1992 by Jacqueline A. Homan
ACKNOWLEDGMENTS
I am indebted to many individuals for their contributions to the inception
and completion of this study. My committee, consisting of J. M. Burns,
H. F. Janssen, F. L Rose, M. R. Willig, and J. K. Jones, Jr., assisted by critically
reviewing this manuscript, as well as providing advice on technical questions.
For the loan of traps, I am indebted to R. J. Baker. I thank P. A. Doris for
patience while teaching me radioimmunoassay techniques and for the use of
his laboratory and equipment. For field assistance, I thank P. A. Coffey,
D. L Hall, H. F. Janssen, J. S. Milner, T. R. Mollhagen, G. Palomino,
R. D. Stephens, L L Torres, and W. S. York. J. M. Burns, J. A. Carr, D. L Hall,
M. Maltbie, J. S. Milner, G. Palomino, and L L Torres assisted with laboratory
procedures. Advice and/or equipment was freely given by R. J. Baker,
S. Bilimorla , R. L Blanton, N. L Collie, L D. Densmore, A. Hinko, J. Hubbard,
J. B. Lombardini, T. R. Mollhagen, R. Patino, B. Pence, M. San Francisco, and
T. E. Tennor, Jr. C. D. Williams provided the figures and photographic
documentation. I wish also to thank the landowners who generously allowed
me to trap on their property: W. Stanford, C. Rose, J. F. Daugherty, and
V. Bozeman. For their time and help with statistical analyses, I thank
M. R. Willig, and D. Hubbard; R. Huber wrote the Games and Howell program.
D. L Bustamantes and L E. Tanner aided in preparation of the final manuscript.
TABLE OF CONTENTS
ACKNOWLEDGMENTS
ii
ABSTRACT
iv
LIST OF TABLES
vi
LIST OF FIGURES
vii
CHAPTER
L INTRODUCTION
1
N. MATERIALS AND METHODS
6
Animal Collection
6
Animal Facilities
7
Treatment Groups
8
Surgical Procedures
8
Assays
10
Statistical Analysis
13
in. RESULTS
15
IV. DISCUSSION
29
LITERATURE CITED
44
APPENDIX
53
ABSTRACT
Few studies of desert rodents (Mammalia: Heteromyidae) have been
designed to address the adaptive role of arginine vasopressin (AVP). The
objectives of this study were to determine the effects of water stress on plasma
and pituitary arginine vasopressin (AVP) and the Impact of increased hormone
levels on blood and urine parameters. Adult males of two sympatric species of
heteromyid rodents iOipodomys ordii and Chaetodipus hispidus) were livetrapped on the Llano Estacado of western Texas and acclimated to laboratory
conditions. Experimental groups, which were fed different diets for 15 days,
included: water control (grain, carrots, and water); baseline control (grain and
carrots); and water-stressed (grain only). Plasma and whole pituitary AVP
concentrations, urine and plasma osmolalities, hematocrit, and weight changes
were each compared among experimental groups. Kidneys were processed
for AVP receptor studies. Two-way (species and treatments) analysis of
variance for each variable revealed that the two species reacted differently to
the treatments. Water-stressed C. hispidus had significantly greater
concentrations of pituitary AVP (1.017 ^g AVP/mg) compared to concentrations
in D. ordii (0.088 jig AVP/mg). Dipodomys ordii had higher plasma AVP levels
compared to those in C. hispidus. Urine osmolality increased significantly with
dehydration in both species, but no significant difference existed between
species. Values ranged from 1878 mOsm/kg In water controls to 3535
mOsm/kg In water-stressed C. hispidus, and from 1158 mOsm/kg to
Iv
3267 mOsm/kg for the same treatments in D. ordii. Compared to both control
groups, values for plasma osmolality were higher in water-stressed D. ordii but
lower in 0. hispidus. Hematocrit increased in both species with dehydration.
Limited data indicated that receptor concentration in all treatment groups was
higher for C. hispidus than for D. ordii. Together, data indicate that the aridadapted D. ordii appears to be more stressed by 15 days of dehydration than
the more mesic-adapted C. hispidus.
LIST OF TABLES
1.
Results of two-way ANOVA between species and treatments
21
2.
Dissociation constants and maximum binding sites for renal AVP
receptors
22
3.
Plasma AVP values reported for different species (pg/ml)
43
A-1
Means ± standard errors for pituitary AVP
54
A-2
Means ± standard errors for plasma AVP
55
A-3
Means ± standard errors for urine osmolality
56
A-4
Means ± standard errors for plasma osmolality
57
A-5
Means ± standard errors for hematocrit
58
A-6
Means ± standard errors for weight change
59
A-7
Means ± standard errors for total pituitary AVP
60
VI
LIST OF FIGURES
1.
Means ± standard errors for pituitary AVP
23
2.
Means ± standard errors for plasma AVP
24
3.
Means ± standard errors for urine osmolality
25
4.
Means ± standard errors for plasma osmolality
26
5.
Means ± standard errors for hematocrit
27
6.
Means ± standard errors for weight change
28
VII
CHAPTER I
INTRODUCTION
Heteromyids are predominantly a family of arid-adapted, almost
exclusively North American rodents, the ancestors of which date from the
Oligocene (Wood, 1935; Hafner and Hafner, 1983). Extant genera include
pocket mice iPerognathus, Chaetodipus), kangaroo mice iMicrodipodops),
kangaroo rats (Dipodomys), and spiny pocket mice iUomys, Heteromys).
Members of the family possess an array of adaptations, including both
behavioral and physiological traits, that allow them to flourish in the arid regions
of the Western Hemisphere from southern Canada to northern South America
(Reichman and Brown, 1983).
These mammals are nocturnal, burrowing, and primarily granivorous, and
all possess external, fur-lined cheek pouches. Some species appear to require
green vegetation at least at some time in their life cycle (Kenagy, 1972, 1973;
Chew and Mitchell, 1965; Frank, 1988a, 1988b). Insects, other arthropods, and
fungi are included in the diets of kangaroo rats and pocket mice (Blair, 1937;
Johnson, 1961; Flake, 1963; Alcoze and Zimmerman, 1973; O'Connell, 1979).
Heteromyids are generally believed to be independent of free water.
Water conservation by heteromyids is well-developed, but not to the
same extent in all genera (Hudson and Rummel, 1966; Foreman and Phillips,
1988). Some species in the genera Chaetodipus, Perognathus, Microdipodops,
and Dipodomys go into torpor when food is unavailable or when exposed to
temperatures below a certain minimum, thus temporarily decreasing the
necessity for water regulation (Bartholomew and Cade, 1957; Morrison and
Ryser, 1962; Brown and Bartholomew, 1969; Yousef and Dil, 1971; Kenagy,
1973). Respiratory water loss is believed to be minimized In some species by
the complex structure of the nasal passages (Jackson and Schmidt-Nielsen,
1964), by microhabitat selection (Lindborg, 1952; Nichter, 1957). by behavior
(Kenagy, 1976), and by maintenance of high burrow humidity (Schmidt-Nielsen
and Schmidt-Nielsen, 1950; Kay and Whitford, 1978).
Water retention by the kidney Is another means whereby these animals
conserve water. Kidney morphology of heteromyid differs from that found in
nondesert rodents in that about 30% of the nephrons are juxtamedullary; these
nephrons have long loops of Henle, some of which extend into the papilla
(Altschuler et al., 1969). This configuration enhances water retention (Jamison,
1987). The kidneys of some heteromyids also show an elongation of the inner
and outer medulla, which augments urine concentrating ability and, therefore,
conserves water (Jamison, 1987; Lawlor and Geluso, 1986). That these
morphological adaptations perform well is evidenced by high urine
concentrations achieved by members of this family. Indeed, heteromyids
concentrate urine to a greater extent than many other mammals (Randle and
Haines, 1976; Nagle et al., 1981; Wright et al., 1977; Hudson and Rummel,
1966; Schmidt-Nielsen and O'Dell, 1961).
Other essential features of urine concentrating ability and water
conservation are the production of arginine vasopressin (AVP) in the
hypothalamic paraventricular (PVN) and supraoptic nuclei (SON), and Its
secretion from the neurohypophysis. Arginine vasopressin Is an octapeptide
hormone that acts via cell surface receptors. It affects specific receptors (V2)
by activating adenylate cyclase as a second messenger in the renal medullary
collecting ducts, rendering them more permeable to water. In general, high
concentrations of AVP stimulated by increased plasma osmolality or by volume
depletion suggest a greater ability to reabsorb water (Dunn et al., 1973;
Jamison et al., 1985; Ausiello et al., 1987; Hays et al., 1987).
Few studies have been conducted that addressed the effects of
dehydration and AVP concentration in heteromyids. In D. merriami, Ames and
van Dyke (1950, 1952) found high concentrations of this hormone in plasma,
urine, and the pituitary compared to levels found in the laboratory rat (Rattus
norvegicus). The concentration of plasma AVP in water-stressed D. spectabilis
is several times higher than in laboratory rats (Stallone and Braun, 1988). The
effects of dehydration induced by hypertonic saline Injection on putative stores
of AVP In the neurohypophysis have been investigated by Scott (1969);
hormone stores are quickly depleted then replenished after treatment. Hatton
et al. (1972) found that xeric-adapted rodents, regardless of family, had greater
numbers of cells in the SON and a large proportion of these cells had multiple
nucleoli. Increased number of nucleoli in AVP-producIng cells was correlated
with an increase In production of the hormone in desert rodents.
Little is known about the effects of water stress on heteromyids with
respect to the consequences of chronically high circulating levels of AVP. In
many mammals, tissues subjected to high protein hormone levels respond by
decreasing the number of receptors. This does not diminish the hormone's
response because of the presence of "spare" receptors (Baddouri et al., 1984;
Steiner and Phillips, 1988). Even though AVP functions in water reabsorption
from renal collecting tubules, this alone may not be enough to maintain
homeostasis with regard to water balance In xeric-adapted mammals.
The purpose of this study was to measure the effects of dehydration on
plasma and pituitary AVP levels, renal AVP receptor levels, urine and plasma
osmolality, hematocrit, and weight loss in two sympatric heteromyid rodents.
Both species are challenged by the same environmental pressures; thus a
study of the two might reveal contrasts or parallels, or both, in adaptations.
Dipodomys ordii Woodhouse, 1853 and Chaetodipus hispidus (Baird, 1858)
were chosen for this study because they are locally abundant, occur in the
same general habitat, are of approximately the same size, and are easily
captured and kept in captivity. Dipodomys ordii, Ord's kangaroo rat, is a
mostly bipedal rodent of moderate size that inhabits open, sandy areas
(Garrison and Best, 1990), and is considered to be a true desert rodent (Mares,
1983). This kangaroo rat is widely distributed from southern Hildago, Mexico,
north through western Texas, Oklahoma, Kansas, and the Dakotas, to southern
Alberta and Saskatchewan and northwest to southern Oregon (Hall, 1981).
Chaetodipus hispidus, the hispid pocket mouse, is quadrupedal and occupies
more dense vegetation on heavier soils than kangaroo rats (Paulson, 1988). It
ranges from southern North Dakota through central Mexico, and from extreme
western Louisiana to southeastern Arizona (Hall, 1981).
CHAPTER II
MATERIAL AND METHODS
Animal Collection
To prevent reproductive hormones from confounding the effects of
dehydration, only males of both Dipodomys ordii and Chaetodipus hispidus
were utilized (Howe and Jewell, 1959). All animals were caught in Sherman live
traps baited with dry rolled oats between May and October, 1991. In cold
weather, a cotton nest was provided. Traps were set just prior to sunset and
checked at sunrise; they were moved every two to four days after males were
trapped out of an area. When caught, animals were identified as to species
and sex. Juveniles and females were released immediately; males were
transported to the animal holding facility in the traps in which they were
captured.
Three Texas collecting sites were used: Lynn Co., 2 mi S, 4 mi W Slide;
Lubbock Co., 1.5 mi W Slide; and Lamb Co., 3.8 mi N Fieldton. The first two
localities occur within the same contiguous four sections of Conservation
Reserve Program (CRP) land and were planted primarily in weeping love grass
(Hall, 1991). Both were characterized by terraces and small bare patches
between growths of dense vegetation, which included love grass, Johnson
grass, and annuals. The Lamb County site was on an arm of the Muleshoe
Sandhills, characterized by sandy soil with large patches of bare, loose sand
between clumps of vegetation, which included mesqulte, catclaw, yucca, and
mixed grasses. (See Pesaturo et al., 1990 for a detailed description of the
area.) Rodent burrows were conspicuous at all localities. A total of 10,000 trap
nights was required to obtain a sufficient number of animals for this study.
Voucher specimens were deposited in The Museum, Texas Tech University,
Lubbock, Texas.
Animal Facilities
Rodents brought to the laboratory were transferred to standard plastic
laboratory animal cages with metal wire tops. Cages for Dipodomys measured
25 X 43 X 20 cm and those for Chaetodipus v^ere 20 x 25 x 15 cm. Each
animal was provided with dry sand on the bottom of the cage, an empty metal
can for an artificial burrow, and cotton for bedding (or plugging the "burrow").
Food, provided ad libitum, consisted of wild birdseed containing sunflower
seed, millet, and sorghum (no more than 10% protein and 4% fat). All animals
were given slices of carrot every other day. The light/dark cycle was 12/12
(light from 6:00 AM to 6:00 PM CST), and the temperature was a constant
22° C. Animals were allowed to acclimate for at least four weeks before being
assigned to a treatment group. Protocol approval for this study was given by
Texas Tech University Animal Care and Use Committee #91221.
Treatment Groups
After acclimation, 20 males of each species were assigned to one of
three treatment groups. Preliminary experiments were performed to determine
the optimum diet and time course for treatments. Fifteen days of a mixed grain
diet caused significant elevation in plasma AVP and urine concentrations
without causing undue harm to the animals. Each group was maintained on a
particular treatment diet for 15 days. Baseline control (B) animals were allowed
to continue on the maintenance diet of free-choice grain and carrots every other
day. Hydrated controls (W) were treated the same as baseline controls, except
they were provided with free access to distilled water. The water-stressed (S)
group received ad libitum grain but no water and no succulent material. All
animals (except baseline controls) were weighed and assigned an individual
number at the beginning of the treatment period. Treatment groups were
created on a staggered schedule so that only one group (20 animals) was
killed per day. All animals were processed between 16 and 30 November,
1991.
Surgical Procedures
On the day of an experiment, all 20 animals of one species in a treatment
group were moved from the animal facility to a laboratory where they were left
undisturbed for at least one hour before processing began. Animals were
removed from cages, injected intraperitoneally with sodium pentobarbital
8
(50 mg/kg) and weighed. After an animal had reached stage III of anesthesia, it
was placed on Its back and a mid-ventral incision was made from the urethra
through the thorax. Cardiac puncture with a heparinized 22-gauge needle was
used to obtain about 1.0 ml of blood. This procedure took approximately 10-15
seconds. The heart was then severed from the great vessels. Blood was
placed in heparinized tubes on ice; a small sample was taken for determination
of hematocrit. Tubes were spun at 1400 x g for 10 min at 4° C on a Beckman
T-J6 centrifuge. Plasma was removed and stored in 1.5-ml microfuge tubes on
ice until osmolalities were determined, after which they were stored at -20" C.
Urine was aspirated from the bladder with a 26-gauge, 5/8-inch needle attached
to a 1.0-cc syringe. Urine was immediately diluted and stored in 0.5-ml
microcentrifuge tubes on ice. Kidneys were removed and placed in
homogenization buffer (see Receptor Preparation) on ice until all animals were
processed. The head was removed and placed on ice. The calvarium was
opened, and the brain retracted to expose the pituitary gland. After retracting
the basal membrane, the entire pituitary gland was removed and placed in 0.5
ml of 1 N acetic acid (Mlaskowski et al., 1988). Each gland was sonicated for
3-6 seconds on a model W-225R Heat Systems-Ultrasonics sonicator/cell
disrupter. Tubes were spun for 30 min in a Herme centrifuge at 11,000 x g at
4° C. All but 0.1 ml of supernatant was removed for use in the
radioimmunoassay for AVP.
Both plasma and urine osmolalities were measured on a Wescor model
5100C vapor pressure osmometer. Urine was diluted 1:10 in millipore-filtered
water and osmolality was determined within 24 hours of sampling. Plasma
osmolality (mOsm/kg) was measured in undiluted samples within four hours of
withdrawal from animals. All values used in data analyses represent means of
triplicate readings.
A small blood sample was taken in 75-mm capillary tubes and spun on a
model MB International micro-capillary centrifuge for two min. Hematocrit was
read as a per cent on a micro-hematocrit capillary tube reader.
Assays
Radioimmunoassay. Extraction and radioimmunoassay (RIA) procedures
for vasopressin followed those published elsewhere (Doris, 1988). Thawed
plasma samples were spun for 15 min at 1400 x g on a Beckman J-B6
centrifuge. An aliquot of 0.25 ml was taken for extraction on Analytichem C-18
columns. Utilizing a Vac-Elute system (Analytichem), columns were prewetted
with 1.0 ml of absolute methanol, rinsed with 2.0 ml of distilled water, and 0.25
ml of sample was added. The column was rinsed with Tris-base and eluted
with 1.0 ml of methanol:water:formic acid (80:18:2). The eluent then was dried
under a stream of air. Sample tubes were capped and frozen at -20'* C until
the day of assay. Pituitary samples were thawed and extracted in the same
manner but without centrifugation. Plasma and pituitary extracts were
10
reconstituted In 0.5-ml assay buffer containing 290 mM Tris-HCI, 0.1 % sodium
azide, and 0.2 % normal rabbit serum. The RIA for the hormone was
performed as follows: Duplicate standards of 0.19-100 fm AVP or unknowns of
100 ^1 were incubated with antiserum (donated by P. A. Doris) for 24 hours at
4** C. The final concentration of the antiserum was 1:60,000 in an incubation
volume of 0.5 ml. lodinated AVP (New England Nuclear, specific activity 2200
Ci/mmol) then was added In the amount of 10,000 cpm per tube and incubated
24 hours at 4** C. A second antibody, goat-antirabbit gamma globulin (10 \i\).
and polyethylene glycol (100 \i\, 25% w/w) were used to separate bound from
free hormone. Tubes were then centrifuged at 1800 x g for 15 min; the liquid
was aspirated and pellets were counted in a Beckman 5500 gamma counter
equipped with a DP5500 Data Reduction System.
Receptor Preparation. The AVP receptor purification was conducted
using a modification of Hinko and Rapp (1989) and Soloff et al. (1989).
Kidneys were kept on iced homogenization buffer [10 mM HEPES,
1 mM EDTA, 0.01% bacitracin, 250 mM sucrose, 1 mM benzamidine, 4 mM
sodium azide, and 0.2 M phenylmethylsulfonyl fluoride]. After all 20 animals in
a group were processed, AVP receptor purification was begun. All steps were
performed on ice in 20 volumes of buffer. Fat was removed and kidneys were
decapsulated. The cortex was separated from the medulla under a dissecting
microscope. Cortical tissue pooled from all 20 animals in a treatment group
was finely minced with a scalpel or razor blade, and homogenized with a teflon
11
pestle in a glass tube. The homogenate wasfilteredon a thin layer of glass
wool and spun at 1000 x g for 5 min. at 4* C on a Beckman J2-21 centrifuge.
The supernatant was layered in Beckman ultraclear centrifuge tubes on a
gradient consisting of 0.4 M sucrose and 1.0 M sucrose dissolved In the
homogenization buffer. Tubes were spun at 111,000 x g for 1.5 hours at 4** C
in a Beckman L8-70 ultracentrifuge. The cloudy layer containing the membrane
was transferred into clean ultraclear tubes which contained ice cold dilution
buffer (100 mM Tris-HCI, 10 mM MgCI2, 0.01% bacitracin, 1 mM benzamidine. 4
mM sodium azide, and 0.2 M PMSF). Tubes were spun at 106,000 x g for 30
min at 4° C. Supernatant was removed and membranes were resuspended in
1 ml ice cold dilution buffer and vigorously vortexed, then sonicated for 2-4 sec
to disperse membrane fragments. Protein concentration was determined by
the Bradford method (Bradford, 1976). Membrane fractions were allquoted into
1.5 ml microfuge tubes, sealed and frozen at -80° C until used in receptor
assay.
Receptor Assay. On the day of assay, membranes were thawed on ice.
Fractions containing 300 \iQ protein were incubated with of ^H-AVP (New
England Nuclear, specific activity 66.7 Ci/mmol) ranging from 0.1 nM to 2.4 nM.
Nonspecific binding was determined with the addition of 1000-fold unlabeled
AVP. Incubation in a final volume of 300 ^1 was carried out at 22° C for 45 min.
Reactions were stopped by pipetting 250 ^1 of the mixture on to Whatman
GS/C glassfilterspresoaked in assay buffer and 0.3% (w/w) polyethylenimine
12
(Hinko and Rapp, 1989). Filters then were rinsed with 15 ml of assay buffer
and placed in 20-ml polyethylene scintillation vials containing 10 ml of Ecolume
(ICN). After 24 hours, samples were counted for 10 min on a Beckman LS7500
liquid scintillation counter. Each treatment group from each species was
assayed separately. Because the amount of membrane was limited, only one
assay for each treatment group was possible. Linear regression analyses for
receptor concentration (Bmax) and dissociation constant (Kd) were carried out
by the EBDA method (McPherson, 1985) included in the RADLIG group of
computer programs (McPherson, 1986).
Statistical Analysis
Considerable individual variation characterized most of the data.
Therefore, outliers were determined and eliminated by Biostat I (Pimentel and
Smith, 1986). Differences in parameters (urine osmolality, plasma osmolality,
plasma AVP, pituitary AVP, hematocrit and weight loss) between species and
among treatment groups were determined by separate two-way analysis of
variance (ANOVA) performed using SPSS-X (SPSS, Inc., 1988). Cochran's C
test was used to evaluate equality of variances. Since variances were
heteroscedastic. pair-wise comparisons of means were tested using the Games
and Howell method (Sokal and Rolf, 1981). Analysis of covariance (ANCOVA)
was performed with BMDP (Dixon and Brown, 1979). Significance was
13
recognized at three levels: significant (0.05 ^ P > 0.01), highly significant
(0.01 k P > 0.001), and very highly significant (P ^ 0.001).
14
CHAPTER I
RESULTS
Captive kangaroo rats occasionally were seen drinking from water
bottles and many readily consumed carrots when they were offered. In
contrast, pocket mice were not seen drinking and often rejected carrots or
buried them immediately. Although measurements were not taken, it was
apparent that water levels in bottles decreased more quickly in kangaroo rat
cages than in pocket mouse cages. Most pocket mice plugged metal cans
with cotton. In contrast, kangaroo rats spread the bedding material around the
cages; some used the can only for a retreat when disturbed and slept on a
loose pile of cotton bedding.
Individuals of both species responded to anesthetic within 3-5 min. The
actual process of drawing blood and killing each animal was completed in
about 30 seconds. All but one specimen was considered to be healthy at the
end of the treatment period; one kangaroo rat In the water-stressed group was
in poor condition and was not used in the study. No animals displayed skin
tenting, an indication of severe dehydration.
Results of the two-way ANOVA between species and among treatment
groups for each parameter are shown in Table 1. Except for urine osmolality
(NS, P = .858) and hematocrit (NS, P = 0.055). differences between species for
parameters were very highly significant. All treatment effects were either
15
significant (pituitary and plasma AVP) or very highly significant (hematocrit,
plasma osmolality, urine osmolality, and weight change). Interaction between
species and treatment was very highly significant for all physiological
parameters except hematocrit (NS) and weight change (highly significant). A
significant interaction demonstrates that the two species responded differently
to treatments.
Inasmuch as Cochran's C test revealed heteroscedascity for all
physiological parameters, statistical differences between treatments within
species, as well as those between species within treatments, were evaluated by
the Games and Howell test (Sokal and Rolf, 1981). These results are displayed
In Figures 1-6. Although this a posteriori test Is less powerful than ANOVA, it is
considered to be a true measure of statistical differences, and holds
experimentwise error rate at 0.05. Treatment categories presented in figures
are arranged so that a progression from the most hydrated state (water-control)
to the most dehydrated (water-stressed) appears from left to right. Descriptive
statistics for all combinations of species and treatments appear as separate
tables for each physiological parameter in the Appendix.
Pituitary AVP (jig AVP/mg pituitary) concentrations were significantly
different for all three treatment groups between species (Fig. 1). The mean
pituitary weight was 0.528 g for C. hispidus, whereas that for D. ordii was
approximately five times larger (2.644 g). Consequently, while the whole
pituitary content of AVP in one D. ordll v/as actually greater than that for one
16
C. hispidus (Table A-7), division of hormone amount by the pituitary weight
equalized the quantity of hormone produced by each species. Note that mean
pituitary AVP levels (Fig. 1) for C. hispidus v/as not significantly different among
treatment groups (W, 0.975 jig AVP/mg; B, 0.924 \LQ AVP/mg; S, 1.017 jig
AVP/mg). but that the water-stressed mean for D. ordll (0.088 jig AVP/mg) was
significantly different from all other groups (W. 0.434 jig AVP/mg; B, 0.442 ^g
AVP/mg).
Figure 2 shows mean plasma AVP concentrations measured in
picograms/mllliliter (pg/ml). In the kangaroo rat, mean AVP levels decreased
with water stress from 1378 pg/ml (W) to 818 pg/ml (S), but in C. hispidus, the
mean AVP levels increased from 423 pg/ml to 578 pg/ml. These opposite
trends were shown as a very highly significant interaction for this parameter
(Table 1). Water-stressed animals displayed no mean differences between
species ( C hispidus, 578 pg/ml; D. ordll, 818 pg/ml), but both baseline and
water-control treatments produced in significant mean differences between
species (the latter 423 pg/ml in C. hispidus, 1378 pg/ml in D. ordl'r, the former
469 pg/ml in 0. hispidus, 1010 pg/ml in D. ordll). Mean plasma AVP (Fig. 2) in
D. ordll was far greater than in C. hispidus, possibly because the former had a
larger pituitary and produced more hormone per ml of blood. The patterns for
AVP content in both plasma (Fig. 2) and pituitary (Fig. 1) across treatment
groups were the same.
17
Results for urine osmolality are displayed in Figure 3. Only baseline
control means were significantly different between species (C. hispidus, 1924
mOsm; D. ordll, 2830 mOsm). Within species, mean values for water-stressed
C. hispidus (3535 mOsm) were significantly higher than those in the other two
treatments (W, 1878 mOsm; B, 1924 mOsm). For water-stressed D. ordll, the
mean for urine osmolality (3267 mOsm) was statistically indistinguishable from
baseline controls (2830 mOsm). but was significantly higher than in the watercontrol group (1158 mOsm).
The two control treatments affected the two species differently with
respect to plasma osmolality (Fig. 4). Within D. ordll, no treatment had a
significant effect on plasma osmolality; in C. hispidus, baseline control means
(326 mOsm) were significantly higher than water-stressed means (310 mOsm).
but neither of these groups were statistically different from water controls (320
mOsm). For C. hispidus, plasma osmolality means were higher in both of these
treatments (W: C. hispidus, 320 mOsm; D. ordll, 308 mOsm; B: C. hispidus,
326 mOsm; D. ordll, 314 mOsm).
Hematocrits of both species responded to greater dehydration in a
similar way, increasing from means of 45.2% (W) to 50.0% (S) in C. hispidus
and from 44.5% (W) to 48.5% (S) in D. ordll (Fig. 5). No significant mean
differences existed between species within any of the treatment groups. Within
species, water-control and water-stressed means were significantly different
18
from each other in both D. ordll (W, 44.5%; S. 48.5%) and C. hispidus
(W, 45.2%; S. 50.0%).
Changes in body weight between water control and water stress
treatments are shown in Figure 6. Water-stressed D. ordll lost significantly
more weight (-7.33 g or 10.2%) over the course of the experiment than did
C. hispidus within the same treatment group (-1.73 g or 3.4 %). Differences in
weight gain (+ 0.44 g in C. hispidus) and weight loss (-0.88 g in D. ordll)
among water-control animals were not significant.
Table 2 shows dissociation constants (KQ) and receptor concentration
(BR/IAX)
for treatment groups and species. These data represent results of one
experiment for each treatment group and each species. The amount of kidney
tissue available was limited; the mean weight of one kidney was 0.180 g for
C. hispidus and 0.331 g for D. ordll. The amount of receptor (protein) obtained
was approximately 5000 jig per 40 kidneys in each C. hispidus group and 5400
^g in D. ordll groups. The dissociation constants in both species within the
water-control treatment were higher than might have been expected; affinity
between hormone and receptor is not thought to change with state of
hydration. The affinity (I/KQ) between AVP and Its receptor was greatest in
baseline control groups for both species, and the greatest concentrations of
binding sites (BMAX) were in the water-control groups.
Analysis of covariance for hematocrit, plasma osmolality, and urine
osmolality, performed with plasma AVP as the covariate demonstrated that only
19
urine osmolality and AVP in C. hispidus showed a significant relationship
(P = .037). Moreover, the effect of AVP on urine osmolality was consistent
within each of the three treatments. A posteriori tests indicated that baseline
control means for C. hispidus were not different from water-control means, but
that the latter group was very highly significantly different from the waterstressed group; baseline control means were very highly significantly different
from those in the water-stressed group.
20
:?
a.
O
O
o
v
V
V
O)
CVI
CO
CM
O)
CM
lO
CO
(O
O)
GO
GO
in
^_
^
GO
^
LU *d
z3
CM
2
CO
d
t
o
o
V
V
Oi
CM
y—
,^_
o
o
v
,,_
O
oV
.ti
O
O
(0
E
a>
d"
CO
CO
O
A
o
o
II
63.7
CO
O
O
13.6
BOD
WEIG
CHAN
OL
II
-*
CL
'^
^
(0
<D
C3)
5"
E
C
,_
o
o
(0 - ^
Q.
"
• ^ JZ.
T3
g)
V
d §
o
(D
E
c
(0
M
O
'o
O
Q.
(0
c
•ft,,
ti
cc
o
O
^
CO
s
:?
LU
0<
0)
o
•
ii
QL
CI
CO
ZE
Q.
I
O
CO
in
id
00
d
LO
in
.,_
(O
CM
GO
o
""i-
CO
CM
CO
O)
Oi
'^
T—
.,_
^_
CM
O
.,_
I-
o
o
-
CM
£
O
o
o
v
,^_
o
o
;<Q-
V
b
Q.
3 iS
o
o
o
^
3
O
(0
GO
x;
<]>
(1>
CM
98.4
0)
h^
CO
C7>
V
in
GO
V
• ^
"^
GO
CM
,,_
q
qV
—
.2
.^
X
(0
O
CO
-
(0
>
E E
(0
o
T3
o
<
s
O)
CM
in
• ^
o
o
o
o
T—
WUJ
O
.2
i3
(0
LU
Q_
CO
CVJ
CM
~^
hX
O
(0
X
lii
21
X
.
o
X
CO
CO
2 s s
•D T3 73
( D O C )
•c r -c
o o
E o
Q. Q. Q.
0)
k_
i_
W CL ^
<D •»— (D
(D T3 i 3
^
k-
0)
.Q
(D
i3
w_
C3) CM
<1>
Q ^
O
0)
k_
E E E
3
C4
T»
TREATMENT
T3
.
„ o o
0)
o
CO
.
5
3
<]>
^ 8
free
las
c
CO
12.7
c»
to
11.7
CO J
3
3
Table 2.
Dissociation constants and maximum binding sites for
renal AVP receptors. Values represent results from one
experiment performed on tissue pooled from 20 animals
in each group.
SPECIES
TREATMENT
nmol/mg
nM
C hispidus
WATER CONTROL
18.0
1.65
BASELINE
2.6
0.60
6.6
0.78
3200.0
147.15
1.3
0.07
2.8
0.29
CONTROL
WATER STRESS
D. ordii
WATER CONTROL
BASELINE
CONTROL
WATER STRESS
22
1.2^
«9
Q.
CD
Q
E
a:
C. hispidus
D. ordll
>
<
Water
Control
(W)
(a)
C. hispidus
(b)
D. ordii
Baseline
Control
(B)
W^s
w
B
Water
Stress
(S)
gWS
3WB
B^
8
Figure 1. Results for pituitary AVP. (a) Means ± standard errors;
asterisks (*) denote significant differences between species within
treatments, (b) In pair-wise comparisons between treatments
within species, common superscript letters show pairs with no
significant differences. Experiment-wise error rate was held
constant at 0.05 via the Games and Howell method.
23
1600-,
14001200
o
Q.
1000
800
^
0. hispidus
gj
D. ordll
600
400-1
200
(a)
(b)
Water
Control
(W)
C. hispidus
W
D. ordii
W
Baseline
Control
(B)
Water
Stress
(S)
88
B
ws
WB
BS
B ws
WB
Figure 2. Resuts for plasma AVP. (a) Means ± standard errors;
asterisks (*) denote significant differences between species within
treatments, (b) In pair-wise comparisons between treatments
within species, common superscript letters show pairs with no
significant differences. Experiment-wise error rate was held
constant at 0.05 via the Games and Howell method.
24
4000-,
3000C3»
E
<n
O
E
2000-
Q
M
C. hispidus
D. ordll
1000-
(a)
Water
Control
(W)
Baseline
Control
(B)
Water
Stress
(S)
C. hispidus W
B^
8
D. ordii
B^
8^
(b)
W
Figure 3. Results for urine osmolality, (a) Means ± standard
errors; asterisks (*) denote significant differences between species
within treatments, (b) In pair-wise comparisons between
treatments within species, common superscript letters show pairs
with no significant differences. Experiment-wise error rate was
held constant at 0.05 via the Games and Howell method.
25
330-.
320.
Ol
310-
E
0)
O
Q
M
300.
E
C. hispidus
D. ordll
290280
(a)
C. hispidus
(b)
D. ordii
Water
O)ntrol
(W)
Baseline
Control
(B)
WBS
B^
8^
gWS
_WB
BS
Water
Stress
(S)
8
W
Figure 4. Results for plasma osmolality, (a) Means ± standard
errors; asterisks (*) denote significant differences between species
within treatments, (b) In pair-wise comparisons between
treatments within species, common superscript letters show pairs
with no significant differences between treatments.
Experiment-wise error rate was held constant at 0.05 via the
Games and Howell method.
26
o
E
o
>
i
Q
m
o
C. hispidus
D. ordll
(0
Q.
Water
Control
(W)
(a)
C. hispidus W
(b)
D. ord//
Baseline
Control
(B)
B
B
w^
B
Water
Stress
(S)
ws
B
ws
B
Figure 5. Results for hematocrit, (a) Means ± standard errors;
asterisks (*) denote significant differences between species, (b) In
pair-wise comparisons between treatments within species,
common superscript letters show pairs with no significant
differences. Experiment-wise error rate was held constant at 0.05
via the Games and Howell method.
27
2-|
0 -
^"^^Laas
v^^\li|ii
N^v^NlJII
-2ifi
E
1
-4
Ijjijjjjjjjjjijijjjjij
•k ^^S
CO
-6-
Q
M
C. hispidus
D. ordll
-810-
Water
Control
(W)
(a)
C. hispidus
(b)
D. ordii
Water
Stress
(S)
W^
8^
W
8
Figure 6. Results for body weight change, (a) Means ± standard
errors; asterisks (*) denote significant differences between species
within treatments, (b) In pair-wise comparisons between
treatments within species, common superscript letters show pairs
with no significant differences. Experiment-wise error rate was
held constant at 0.05 via the Games and Howell method.
28
CHAPTER IV
DISCUSSION
For more than 100 years, heteromyid rodents have been the focus of
studies in systematics, evolution, behavior, ecology, anatomy, and physiology
(Osgood, 1900; Wood, 1935; Setzer, 1949; Hafner and Hafner, 1983;
Eisenberg, 1963; Reichman, 1983; Rosenzweig and Winakur, 1969; Price and
Brown, 1983; Howell and Gersh, 1935; Schmidt-Nielsen, 1979). Despite these
extensive studies, no comprehensive investigations have encompassed all
aspects of the physiology of heteromyids. This may be due, in part, to the
diversity of the family and to the narrow focus of the investigations. However,
many valuable contributions have been made to our understanding of the ability
of these unique animals toflourishin arid regions.
This study focused specifically on the effects of dehydration on blood,
urine, and renal parameters in two heteromyids. Dipodomys ordii is considered
to be a true desert Inhabitant, and C. hispidus is regarded as somewhat better
adapted to mesic conditions. Animals collected from the same local habitats
would be expected to confront similar environmental situations. To survive,
animals must have a daily water input that at least equals water output. Water
sources for these rodents include only moisture contained In food (preformed
water) and formation of water from oxidative processes (metabolic water).
However, water is lost through several routes ~ by the kidneys and intestine.
29
respiration, and evaporation through the integument. Survival in a dry
environment should be predicated on abilities to maximize water Intake and
minimize water loss. Although animals from the same environment could solve
similar problems in similar ways, results from this study indicate that D. ordll
and C hispidus did not respond to experimental manipulation in the same
manner. Both species have developed the ability to concentrate urine to a high
degree; however, the physiological strategies employed to maintain
homeostasis appear to be different. Based on these data, it Is suggested that
their adaptive strategies in their native environment also may differ.
Chaetodipus hispidus (mean body weight, 49 g) had a pituitary one-fifth
(0.528 mg) the weight of the pituitary in the kangaroo rat but was able to
maintain stores of AVP for 15 days of dehydration. Also, the pocket mouse
consistently had greater quantities of pituitary AVP per mg of pituitary tissue
(Fig. 1). However, the kangaroo rat (mean body weight, 71 g) with the larger
pituitary (2.644 mg) exhibited a significant depletion of stored hormone after 15
days of dehydration and had significantly less hormone per mg of pituitary.
When quantities of AVP per pituitary were calculated (rather than AVP per mg
pituitary), D. ordll had approximately twice that of C. hispidus in both control
groups (Table A-7). In contrast, hormone amounts in water-stressed
C. hispidus were twice those of D. ordll. If maintenance of AVP stores was a
useful adaptation for desert rodents, then the response of the pocket mouse to
30
experimental treatment appeared to make it a better candidate to withstand
long-term dehydration.
It has been confirmed that after varying periods of dehydration or salt
loading, the AVP-producing cells of the supraoptic (SON) and paraventricular
nuclei (PVN) in the hypothalamus of laboratory rats increase in size, the number
of nucleoli increase, and the endoplasmic reticulum becomes dilated (EhrhardBornstein et al.. 1990; Jones and Pickering, 1969; Hatton, 1990; Hatton and
Walters, 1973; Morris and Dyball, 1974; Tweedle and Hatton, 1976; Watt, 1970).
These cellular events permit greater AVP production to meet the demands of
dehydration.
Although only a limited number of dehydration studies have been
performed on xeric-adapted species, it seems clear that these animals have the
capacity for greater AVP production and storage than the laboratory rat. Ames
and van Dyke (1950) compared neural lobe content of R. norvegicus to
D. merriami and found that hydrated kangaroo rats had three times more AVP
than did laboratory rats. In another study of D. merriami, Scott (1969) found
that the neurohypophysis became depleted of AVP stores within 10-30 min after
intraperitoneal injection of hypertonic sodium chloride, but that by 60 min after
injection, there was almost 100% repletion of the neurosecretory granules
containing AVP. This accelerated rate of recovery might be an adaptation to
desert environments and may be correlated to the increase in numbers of
31
nucleoli recruited for protein synthesis (Scott, 1969). Hatton et al. (1972)
studied both mesic and xeric animals - heteromyids {D. merriami and
D. spectabilis) and murids - (Peromyscus callfornlcus, P. eremlcus and
Merlones ungulculatus) and concluded that the SON from xerIc animals,
regardless of family, had a greater number of cells and that 20-60% of these
cells had multiple nucleoli. Castel and Abraham (1972) compared
neurohypophyseal AVP in spiny mice (Acomys cahlrinus and A. russatus) with
that in /?. norvegicus and Mus musculus. They found that the laboratory rat
neural lobe depleted its stores of AVP; the other three species showed an Initial
decrease in AVP at 14 days, but demonstrated repletion of the hormone in the
pituitary after 22 days.
In view of other studies involving pituitary stores of AVP, the 15-day time
course followed in this experiment might not have been enough to deplete the
pituitary AVP stores In C. hispidus. It is also possible that the pituitary became
depleted during the 15 days of treatment but by the end of the test period,
could have been replenished. Results here demonstrated that pituitary AVP in
D. ordll became depleted, but whether It would become replenished or further
depleted would have to be tested In other studies in which both shorter and
longer periods of time are employed.
Plasma AVP concentrations were not statistically different in any
treatment groups of 0. hispidus or in D. ordll. However, a very highly
significant interaction existed (Table 1): AVP concentrations tended to decrease
32
from water-control to water-stressed groups in D. ordll but increased across
these same groups in C. hispidus (Fig. 2). The patterns In plasma hormone
paralleled those found in pituitary AVP (Fig. 1). Plasma AVP was twice as high
for D. ordll In both control groups compared to those for C. hispidus. This may
seem inconsistent with pituitary AVP values but this is because pituitary AVP is
expressed in amount of hormone (jig) per milligram of pituitary; when the larger
size of the pituitary in D. ordii Is taken into account, it can be seen why the
plasma AVP concentrations are higher (Table A-7).
Handling the animals or the anesthetic might have altered the AVP
response. The consistent use of these procedures in all groups eliminated this
as a variable unless the species responded to these factors in a different way,
or if the treatment affected the response within a taxon. Additional studies
using other methods of anesthesia or vascular access, or both, could address
this question. Other studies have reported substantial variation In plasma AVP
values even within species (Table 3). A number of factors, including specific
experimental treatments, type of method used to determine AVP concentration,
and Intrinsic differences between species or strains of rats could Induce such
variation.
Urine concentration in D. ordii and C. hispidus Increased with
dehydration. In the baseline control group, C. hispidus, which did not readily
take carrots, there was significantly lower urine osmolality (Fig. 3) than in
33
D. ordll, which often actively took carrots. The similarity of values for urine
concentration in both species in water-stressed groups indicated that the renal
concentrating mechanisms in both species were capable of reabsorbing water
from the medullary collecting ducts upon stimulation by increased plasma
osmolality and increased circulating AVP (Dunn et al., 1973; Yagil and Sladek,
1990). In D. ordll, significant differences between baseline controls and water
controls were possibly the result of water consumption in this species.
Anecdotal and experimental information about dipsogenic behavior in kangaroo
rats abounds, but little is known about this phenomenon in pocket mice (Boice,
1972; Vander Weele, 1975). One study of D. ordll Uom Nebraska suggested
that this species was not water independent but investigators offered no
suggestions for the source of water in the native environment of the animals
(Fairbanks et al., 1983).
Urine osmolality values reported here are comparable to those on record
for other heteromyids (MacMillen and Hinds, 1983; MacMillen and Lee, 1967;
and Hudson and Rummel, 1966). It is apparent from other studies that the
experimental conditions of high temperature and a high protein diet result in the
most concentrated urine. The combination of anatomical specialization in the
kidney (Vimtrup and Schmidt-Nielsen, 1952; Schmidt-Nielsen and O'Dell, 1961;
Altschuler et al., 1969; Lawlor and Geluso, 1986) and greater ability to produce,
secrete, and store AVP are all thought to make the renal concentrating
34
mechanism of desert rodents an effective adaptation for survival in hot, dry
climates (Stevenson, 1987).
It is noteworthy to compare the results of plasma AVP (Fig. 2) with those
for urine osmolality (Fig. 3). Increased AVP levels cause increased water
retention and, therefore, higher urine osmolality (Dunn et al., 1973; Yagil and
Sladek, 1990). A tendency for a decrease in plasma AVP existed across
treatment groups in D. ordll, but urine osmolality became elevated with
increased dehydration. Both plasma AVP and urine osmolality became
elevated in C. hispidus with increased dehydration. Decreased plasma AVP
levels In D. ordll could have been due to decreased pituitary stores (Fig. 1).
The two species showed differences in their plasma osmolalities only
after dehydration (Fig. 4). Following 15 days of dehydration, the plasma
osmolality of water-stressed C. hispidus v/as 310 mOsm, a value significantly
below the baseline control group (326 mOsm). The kangaroo rat, on the other
hand, showed no significant differences in plasma osmolality between any
treatment groups. The decrease in plasma osmolality in pocket mice after 15
days of water stress may have been due to movement of water from the
intracellular compartment or the interstitial space, or both. This phenomenon
has been demonstrated in other desert species (Schmidt-Nielsen, 1979).
However, more evidence is available to show that water movement occurs
across all fluid compartments at the same rate at approximately the same time.
When there is even a small change in plasma osmolality, water is pulled from
35
interstitial and intracellular compartments (Seldin and Giebisch, 1985). One
possibility for the apparent increase in water In the vascular compartment of
dehydrated C. hispidus could have been related to metabolism of fat. Although
not precisely determined, it appeared that this species possessed more body
fat than kangaroo rats. Lower plasma osmolality could have also resulted from
solute excretionft-omthe plasma into the urine. In this study however, no
attempt was made to measure speciflc electrolytes or other compounds that
are known to affect plasma osmolality (Schmidt-Nielsen et al., 1961; Stallone
and Braun, 1988).
The values for plasma osmolality obtained In this experiment were
comparable to those of other heteromyids but were higher than those found In
laboratory rats. Values for the latter ranged from 293.6 mOsm (Dunn et al.,
1973) to 291 mOsm (Summy-Long et al., 1990). Stallone and Braun (1988)
reported that normally hydrated D. spectabilis had a mean plasma osmolality of
308.6 mOsm/kg, and, that after eight days on soybeans and no water the mean
value increased to 329.7 mOsm/kg. The high values for dehydrated animals
may have resulted from the fact that they were fed a high protein diet in that
experiment. High plasma osmolality can be caused by either a decrease in
water In the blood or an increase in solutes. Stallone and Braun (1988)
determined that the "set point' for AVP release in D. spectabilis was a plasma
osmolality of 306.4 mOsm/kg water. No studies of this kind have been
reported for pocket mice.
36
Hematocrit was the only parameter in this experiment in which no
significant interaction occurred between species and treatments (Table 1,
Fig. 5). In other words, both species showed the same pattern of response to
treatments: increased hematocrit with dehydration. Moreover, hematocrit was
significantly different between the water-control and water-stressed groups for
both the kangaroo rat and pocket mouse. No significant differences were
obtained between species within any treatments. Water-control means of
45.2% (C. hispidus) and 44.5% (D. ordll) were higher than those for hydrated
laboratory rats, which ranged from 36.4% (Dunn et al., 1973) to 42.8% (SummyLong et al., 1990). Selander (1964) reported mean hematocrit values of 52.0%
for adult male C. hispidus and 48.0% for adult male D. ordll, but because
measurements were taken predominately from recently trapped animals, and
the state of (de)hydration was not known. Stallone and Braun (1988) reported
normal hematocrit for D. spectabilis as 43.3%, and after eight days of
dehydration, 50.4%. With Increased dehydration, water would be lost from the
vascular compartment (even with an efficient renal concentrating mechanism),
and with water loss, increases in both hematocrit and plasma osmolality would
follow.
One striking result of this experiment was that water-stressed C. hispidus
actually showed a decrease in plasma osmolality when compared to both
control groups, but an increase in hematocrit. It might be expected that if an
animal retains more water, and thus decreases plasma osmolality, a decrease
37
in hematocrit would follow. However, it may be that in response to dehydration,
pocket mice are able to excrete more solutes from plasma into urine,
consequently lowering plasma osmolality. Elevated hematocrit can result fi-om
either a decrease in plasma water (with dehydration) or an Increase in cell
volume. The latter is unlikely under these conditions.
Body weight loss was significantly different between species in the waterstressed groups (Fig. 6). Kangaroo rats lost an average of 10.2% of their
original body mass, whereas pocket mice lost only 3.4%. Both species in all
treatment groups often were observed husking and eating seeds, especially
those of millet and sunflower, so weight loss was probably not a product of
inadequate food intake. D. spectabilis reportedly lost 22.2% of initial mass after
eight days on soybeans (Stallone and Braun, 1988). Some species apparently
stop eating when deprived of water (Schmidt-Nielsen, 1979; Dicker and Nunn,
1957), but heteromyids maintain or gain weight on dry diets (Schmidt-Nielsen
and Schmidt-Nielsen, 1951). Nevertheless, relative humidity was a crucial factor
in maintaining body weight. When fed only pearled barley, D. merriami
maintained or gained weight at relative humidities above 5-15%, but laboratory
rats and hamsters on the same diet lost weight even at 90% relative humidity
(Schmidt-Nielsen and Schmidt-Nielsen, 1951).
Results concerning renal AVP receptors are inconclusive because only
one experiment using pooled tissues was conducted for each species in each
treatment group (Table 2). Downregulation (decrease in number) of AVP
38
receptors was recorded in studies with laboratory rats (Steiner and Phillips,
1988), pigs (Bockaert et al., 1973), and jerboas (J. orlentalls) (Baddouri et al.,
1984; Butlen et al., 1984). However, other workers reported desensitization or
modulation of the AVP-receptor response, including uncoupling of adenylate
cyclase from the hormone-receptor complex rather than decrease In receptor
number (Ausiello et al., 1987). Both V2 (antidiuretic) and VI (vasopressor)
receptors found on vascular smooth muscle and renal interstitial cells respond
to AVP but through different second messengers. Evidence suggests that both
calcium and prostaglandins released by stimulation of VI receptors modify
urinary concentration in several ways, including alterations in blood flow and
solute transport (Ausiello et al., 1987). The effects of VI receptor activation
stimulated by AVP are probably important in desert animals that have high
circulating concentrations of AVP during times of dehydration, but this aspect of
their physiology has not been Investigated.
Dehydration for 15 days is either not enough to severely stress
C. hispidus; alternatively, the hypothalamic-neurohypophysis AVP replenishing
mechanism in this species is perhaps more efficient than that of D. ordii, or
both. Both species were able to concentrate urine to similar degrees, but
kangaroo rats had higher plasma AVP and osmolar concentrations. Pocket
mice had higher hematocrits, but lost less body mass. Taken together, these
data suggest that the two species have different physiological adaptations to
dehydration.
39
Since there are no apparent permanent sources of free water in desert
rodent environments, it must be assumed that animals obtain their water in the
food they eat. Food habit studies, although not comprehensive. Indicate that
D. ordll consumes more water-containing items than does C. hispidus (Blair,
1937; Alcoze and Zimmerman, 1973; O'Connell, 1979). Studies of food storage
habits of some species of Dipodomys also indicate that they select seeds that
are slightly moldy (Rebar and Reichman, 1984; Reichman et al., 1985) and
higher in water content (Frank, 1988a, 1988b). The physiological significance of
this behavior has not been assessed. At least some heteromyids maintain
higher burrow humidity than would be expec^ted in desert environments
(Schmidt-Nielsen and Schmldt-Nellsen, 1950; Nichter, 1957; Kay and Whitford,
1978). Consuming seeds with greater water content would Increase water
intake; decreased respiratory water loss occurs in air with high humidity. These
kinds of behavorlal adaptations in D. ordll may possibly prevent this species
from becoming water-stressed in its natural environment.
The results of this study have provided insight into the ability of two
heteromyids to respond to water stress. Many other questions remain to be
answered. These include possible interactions between anatomical,
physiological and behavioral characteristics that could influence the ability of
these animals to survive in arid environments (Bartholomew and Caswell, 1951;
Nicolai and Bramble, 1983; Wang and Hudson, 1970; Homan and Genoways,
1978). These interactions, which were beyond the scope of the present study,
40
will require (carefully designed and cx^ntrolled investigations. Subtle differences
between species in behavior and morphology, overlcx)ked in many studies, may
greatly influence physiological stress levels in natural environments.
Summary
1. The pituitary gland was 5 times larger in D. ordll than in C. hispidus.
However, pituitary stores of AVP became depleted In the kangaroo rat with 15
days on a dry grain diet. The pocket mouse maintained its pituitary AVP stores
under the same conditions. The depletion of AVP stores in the more xericadapted rodent was most unexpected.
2. Plasma AVP concentrations were not statistically different within
species among treatment groups, but were signific^antly different between
species (except in the water-stressed groups).
3. Urine osmolality increased in both species with dehydration; the only
significant difference between species was in the baseline control groups.
4. Plasma osmolality in C. hispidus was highest in the baseline control
group but decreased significantly in the water-stressed group. These findings
indicated that the pocket mouse was capable of sun^lving very high plasma
osmolalities, and was able to excrete large amounts of solute from plasma in
order to maintain homeostasis. There were no differences among treatments
for D. ordii. Between the two species, plasma osmolality was significantly
different in all but the water-stressed groups.
41
5. Hematocrit increased in both species with dehydration; there were no
significant differences between species for any treatment.
6. Weight change over the 15 day treatment period resulted in a loss of
10.2 % in D. ordii, and 3.4% for C. hispidus In the water-stressed groups.
7. Renal AVP receptor concentration decreased in both species with
dehydration. Affinity between receptor and hormone was lowest in watercx5ntrol groups, but receptor concentration was highest in these groups.
8. Differences in response to the laboratory treatments suggest, that in
their natural environments, these species possibly employ different behavioral
strategies in order to maintain homeostasis.
42
Table 3.
Plasma AVP values reported for different species in pg/ml.
Treatment and assay methods differed in the experiments.
Taxon
Hydrated
[AVP]
pg/ml
Water
Stressed
[AVP]
pg/ml
R. norvegicus
2-3.0
23
Dunn etal., 1973
R. norvegicus
0.21
2.4
Steiner and Phillips, 1988
R. norvegicus
2.9
26.2
Baddouri etal., 1984
Acomys
-
>300
Castel etal., 1974 In
Baddouri etal., 1984
J. jaculus
-
400-1600
J. orientaJis
76
372
G. gerblllus
-
250-1800
.
120.0
Ames and van Dyke, 1952
6
69
Stallone and Braun, 1988
Reference
MURIDAE
El Husseini and Haggag, 1974
Baddouri etal., 1984
El Husseini and Haggag, 1974
HETEROMYIDAE
D, merriami
p. spectabilis
43
LITERATURE CITED
Alcx)ze, T. M. and E. G. Zimmerman (1973). Food habits and dietary overlap of
two heteromyid rodents from the mesqulte plains of Texas. J. Mamm.
54:900-908.
Altschuler, E. M., R. B. Nagle, E. J. Braun, S. L Lindstedt, and P. H
Krutzsch (1979). Morphological study of the desert heteromyid kidney
with emphasis on the genus Perognathus. Anat. Rec. 194:461-468.
Ames, R. G. and H. B. van Dyke (1950). Antidiuretic hormone in the urine and
pituitary of the kangaroo rat. Proc. Soc. Exp. Biol. Med.
75:417-420.
Ames, R. G. and H. B. van Dyke (1952). Antidiuretic hormone in the serum or
plasma of rats. Endocrinology 50:350-360.
Ausiello, D. A., K. L Skorecki, A. S. Verkman, and J. V. Bonventre (1987).
Vasopressin signaling in kidney cells. Kid. Int. 31:521-529.
Baddouri, K., D. Butlen, M. Imbert-Teboul, F. Le Bouffant, J. Marchetti, D.
Chabardes, and F. Morel (1984). Plasma antidiuretic hormone levels and
kidney responsiveness to vasopressin in the Jerboa, Jaculus orientalls.
Gen. Comp. Endocr. 54:203-215.
Bartholomew, G. A. and T. J. Cade (1957). Temperature regulation,
hibernation, and aestivation in the little pocket mouse, Perognathus
longlmembrls. J. Mamm. 38:60-72.
Bartholomew, G. A. Jr., and H. H. Caswell, Jr. (1951). Locomotion in kangaroo
rats and its adaptive significance. J. Mamm. 32:155-169.
Blair, W. F. (1937). The burrows and food of the prairie pocket mouse.
J. Mamm. 18:188-191.
Bockaert, J., C. Roy, R. Rajerison, and S. Jard (1973). Specific binding of
[^H] lysine-vasopressin to pig kidney plasma membranes. J. Biol. Chem.
248:5922-5931.
Boice, R. (1972). Water addiction in captive desert rodents. J. Mamm.
53:395-398.
44
Bradford, M. M. (1976). A rapid and sensitive method for the quatitation of
microgram quantities of protein utilizing the principle of protein-dye
binding. Analyt. Blochem. 72:248-254.
Brown, J. H. and G. A. Bartholomew (1969). Periodicity and energetics of
torpor in the kangaroo mouse, Microdipodops pallldus. Ecology
50:705-709.
Butlen, D., K. Baddouri, R. M. Rajerison, G. Gulllon, B. Cantau, and S. Jard
(1984). Plasma antidiuretic hormone levels and liver vasopressin
receptors in the Jerboa, Jaculus orlentalls, and rat. Gen. Comp.
Endocrln. 54:216-229.
Castel, M. and M. Abraham (1972). Effects of a dry diet on the antidiuretic
hormone content of the neurohypophysis in spiny mice as compared to
the albino rat and mouse. Gen. Comp. Endocrln. 19:48-55.
Chew, R. M. and O. G. Mitchell (1965). Responses of xeric-adapted rodents to
waterloading. Proc. Soc. Exp. Biol. Med. 120:336-338.
Dicker, S. E. and J. Nunn (1957). The role of the antidiuretic hormone during
water depravation in rats. J. Physiol. 136:235-248.
Dixon, W. J. and Brown, M. B. (1979). BMDP-79: Biomedical Computer
Programs P-series. Univ. California Press, Berkeley.
Doris, P. A. (1988). Angiotensin II Immunoreactivity in push-pull perfusates
from the paraventricular nucleus. Peptides 9:481 -485.
Dunn, F. L, T. J. Brennan, A. E. Nelson, and G. L Robertson (1973). The
role of blood osmolality and volume in regulating vasopressin secretion
in the rat. J Clin. Investlg. 52:3212-3219.
Eisenberg, J. F. (1963). The behavior of heteromyid rodents. Univ. California.
Publ. Zool. 69A-^^ 4.
El-Husseini, M. and G. Haggag (1974). Antidiuretic hormone and water
consen^atlon In desert rodents. Comp. Blochem. Physiol. 47A:347-350.
Erhart-Bornstein, M, N. A. Thorn, and M. Treiman (1984). Chronic osmotic
stimulation reduces vasopressin but not synaptophysin content in rat
neurohypophysis. Neuroscl. Let 119:122-124.
45
Fairbanks, W. S., D. Greegor, L Staudinger, and E. Bitterbaum (1983). Water
conservation of the kangaroo rat. Trans. Nebraska Acad. Sci. 11:27-30.
Flake, L D. (1973). Food habits of four species of rodents on a short-grass
prairie in Colorado. J. Mamm. 54:636-647.
Forman, G. L and C. J. Phillips (1988). Histological variation in the
proximal colon of heteromyid and cricetid rodents. J. Mamm.
69:144-149.
Frank, C. L (1988a). The influence of moisture content on seed selection by
kangaroo rats. J. Mamm. 69:353-357.
Frank, C. L (1988b). The effects of moldiness level on seed selection by
Dipodomys spectabilis. J Mamm. 69:358-362.
Garrison, T. E. and T. L Best (1990). Dipodomys ordii. Mammalian Species
353:1-10.
Hafner, J. C. and M. S. Hafner (1983). Evolutionary relationships of
heteromyid rodents. Great Basin Nat, Mem. 7:3-29.
Hall, D. L (1991). Species diversity and mammalian succession in
Conservation Reserve Program grasslands. Unpublished Masters
Thesis, Texas Tech University, vii -i- 34 pp.
Hall, E. R. (1981). The Mammals of North Americ^a. John Wiley and Sons,
New York, 2nd ed., 1:1-600 + 90.
Hatton, G. I. (1990). Emerging concepts of structure-function dynamics in
adult brain: the hypothalamo-neurohypophysial system. Prog.
Neuroblol. 34:437-504.
Hatton, G. I., J. I. Johnson, and C. Z Malatesta (1972). Supraoptic nuclei of
rodents adapted for mesic and xeric environments: number of cells,
multiple nucleoli, and their distributions. J. Comp. Neurol.
145:43-59.
Hatton, G. I. and J. K. Walters (1973). Induced multiple nucleoli, nucleolar
margination, and cell size changes in supraoptic neurons during
dehydration and rehydration in the rat. Brain Res. 59:137-154.
Hays, R. M., R. Franki, and G. Ding (1987). Effects of antidiuretic hormone
on the collecting duct. Kid. Int 31:530-537.
46
Hinko, A. and J. P. Rapp (1989). Atrial Natriuretic factor and cyclic AMP
modify protein phosphorylation in renal papillary membranes of Dahl
rats. Pp. 53-65 in Progress in Atrial Peptide Research, Vol. III.
Amer. Soc. Hypertension Symp. Ser., B. M. Brenner and J. H. Laragh,
eds. Raven Press, Ltd., New York.
Homan, J. A. and H. H. Genoways (1978). An analysis of hair structure and
Its phylogenetic implications among heteromyid rodents. J. Mamm.
59:740-760.
Howe, A. and P. A. Jewell (1959). Effects of water deprivation upon the
neurosecretory material of the desert rat (Merlones merlones) compared
with the laboratory rat. J. Endocrln. 18:118-124.
Howell, A. B. and I. Gersh (1935). Consen^ation of water by the rodent
Dipodomys. J. Mamm. 16:1-9.
Hudson, J. W. and J. A. Rummel (1966). Water metabolism and temperature
regulation of the primitive heteromyids, Uomys salvani and Liomys
Irroratus. Ecology 47:345-354.
Jackson, D. C. and K. Schmidt-Nielsen (1964). Countercurrent heat exchange
in the respiratory passages. Proc. Nat Acad. Sci. 51:1192-1197.
Jamison, R. L (1987). Short and long loop nephrons. Kid. Int. 31:597-605.
Jamison, R. L, B. Zimmerhackl, and C. R. Robertson (1985). Vasopressin and
the urinary concentrating mechanism: the 1984 perspective. Pp. 485-496
In Vasopressin, R. W. Schrier, ed.. Raven Press, New York.
Johnson, D. R. (1961). Food habits of rodents on rangelands of southern
Idaho. EcTO/opy 42:407-410.
Jones, C. W. and B. T. Pickering (1969). Comparison of the effects of water
deprivation and sodium chloride imbibition on the hormone content of
the neurohypophysis of the rat. J. Physiol. 203:449-458.
Kay, F. R. and W. G. Whitford (1978). The burrow environment of the bannertailed kangaroo rat, Dipodomys spectabilis. In south-central New
Mexico. Amer. Midi. Nat 99:270-279.
Kenagy, G. J. (1972). Saltbush leaves: excision of hypersaline tissue by a
kangaroo rat. Science 178:1094-1096.
47
Kenagy, G. J. (1973). Daily and seasonal patterns of activity and energetics
in a heteromyid rodent community. Ecology 54:^20^-^2^9.
Kenagy, G. J. (1976). The periodicity of daily activity and its seasonal
changes in free-ranging and captive kangaroo rats. Oecologia
24:105-140.
Lawlor, R. M. and K. N. Geluso (1986). Renal structure and body size in
heteromyid rodents. J. Mamm. 67:367-372.
Lindborg, R. G. (1952). Water requirements of certain rodents from xeric and
mesic habitats. Contrlb. Lab. Vert. Biol., Univ. Michigan. 67:1-123.
MacMillen, R. E. and D. S. Hinds (1983). Water regulatory efficiency in
heteromyid rodents: a model and its application. Ecology 64:^52-^64.
MacMillen, R. E. and A. K. Lee (1967). Australian desert mice: independence
of exogenous water. Science 158:383-385.
Mares, M. A. (1983). Desert rodent adaptation and community structure. Great
Basin Nat, Mem. 7:30-43.
McPherson, G. A. (1985). Analysis of radioligand binding experiments: A
collection of computer programs for the IBM PC. J. Pharmacol. Meth.
14: 213-228.
McPherson, G. A. (1986). A Collection of Radioligand Binding Analysis
Programs. Elsevier and BIOSOFT, Cambridge, UK.
Miakowski C , G. L Ong, D. Lukic, and J. Haldar (1988). Immobilization
stress affects oxytocin and vasopressin levels in hypothalamic and
extrahypothalamic sites. Brain Res. 458:137-141.
Morris, J. F. and R. E. J. Dyball (1974). A quantitative study of the
ultrastructural changes in the hypothalamo-neurohypophysial system
during and after experimentally induced hypersecretion. Cell Tiss.
Res. 149:525-535.
Morrison, P. and F. A. Ryser (1962). Hypothermic behavior in the hispid
pocket mouse. J. Mamm. 43:529-533.
48
Nagle, R. B., E. M. Altschuler, D. C. Dobyan, S. Dong, and R. E. Bulger (1981).
The ultrastructure of the thin limbs of Henle In kidneys of the
desert heteromyid (Perognathus penicillatus). Amer. J. Anat
161:33-47.
Nichter, R. (1957). The effect of variation in humidity and water intake on
activity of Dipodomys. J. Mamm. 38:502-512.
Nikolai, J. C. and D. M. Bramble (1983). Morphological structure and
function in desert heteromyid rodents. Great Basin Nat, Mem. 7:44-64.
O'Connell, M. A. (1979). Coexistence of two species of kangaroo rats (genus
Dipodomys) in the Guadalupe Mountains National Park, Texas. Pp. 349371, In Biological investigations in the Guadalupe Mountains National
Park, Texas, H. H. Genoways and R. J. Baker, eds. National Park
Service, Washington, D. C , 442 pp.
Osgood, W. H. (1900). Revision of the pocket mice of the genus Perognathus.
N. Amer. Fauna 18:1-72.
Paulson, D. D. (1988). Chaetodipus hispidus. Mammalian Species, 320:1-4.
Pesaturo, R. J., J. K. Jones, Jr., R. W. Manning, and C. Jones (1990).
Mammals of the Muleshoe Sandhills Bailey, Hale, and Lamb counties,
Texas. Occas. Pap., Mus. Texas Tech Univ. 136:1-32.
Pimentel, R. A. and J. D. Smith (1986). Biostat I. Sigma Soft, 145 pp.
Price, M. V. and J. H. Brown (1983). Patterns of morphology and resource use
in North American desert rodent communities. Great Basin Nat, Mem.
7:117-134.
Randle, H. and H. Haines (1976). Effect of water deprivation on antidiuresis
in Dipodomys spectabilis and Rattus norvegicus. Comp. Blochem.
Physiol. 54A:2^'26.
Rebar, C. and O. J. Reichman (1983). Ingestion of moldy seeds by heteromyid
rodents. J. Mamm. 64:713-715.
Reichman, O. J. (1983). Behavior of desert heteromyids. Great Basin Nat,
Mem. 7:77-90.
Reichman, O. J. and J. H. Brown (1983). Biology of desert rodents. Great
Basin Nat, Mem. 7:1-134.
49
Reichman, O. J., D. T. Wicklow and C. Rebar (1985). Ecological and
mycological characteristics of caches in the mounds of Dipodomys
spectabilis. J. Mamm. 66:643-651.
Rosenzweig, M. L and J. Winakur (1969). Population ecology of desert rodent
communities: habitats and environmental complexity. Ecology
50:558-572.
Schmidt-Nielsen, B. and R. O'Dell (1961). Structure and concentrating
mechanism in the mammalian kidney. Amer. J. Physiol. 200:1119-1124.
Schmidt-Nielsen, B., R. O'Dell, and H. Osaki (1961). Interdependence of urea
and electrolytes in production of a concentrated urine. Amer. J.
P/7ys/o/. 200:1125-1132.
Schmidt-Nielsen, B. and K. Schmidt-Nielsen (1950). Evaporative water loss in
desert rodents in their natural habitat. Ecology 31:75-85.
Schmidt-Nielsen, B. and K. Schmidt-Nielsen (1951). A complete account of the
water metabolism in kangaroo rats and an experimental verification.
J. Cell. Comp. Physiol. 38:165-181.
Schmidt-Nielsen, K. (1979). Desert Animals: Physiological Problems of Heat and
Water. Dover Publ, Inc., New York xiv + 276 pp.
Schmidt-Nielsen, K. and B. Schmidt-Nielsen (1952). Water metabolism of desert
mammals. Physiol. Rev. 32:135-166.
Scott, D. E. (1969). Ultrastructural changes in the neural lobe of the
hypophysis of the kangaroo rat (Dipodomys merriami) following
intraperitoneal injection of hypertonic sodium chloride.
Neruoendocrln. 4:347-367.
Selander, J. A. (1964). The infiuence of body size, season, sex, age and
other factors upon some blood parameters in small mammals.
J. Mamm. 45:598-616.
Seldin, D. W. and G. Giebisch (1985). The Kidney: physiology and
Pathophysiology. Raven Press, New York, 1020 pp.
Setzer, H. W. (1949). Subspeciation in the kangaroo rat, Dipodomys ordii.
Univ. Kansas Publ., Mus. Nat Hist 1:473-573.
50
Sokal, R. R. and F. J. Rohlf (1981). Biometry. 2nd ed. W. H. Freeman Co..
New York, 859 pp.
Soloff, M. S., M. A. Fernstrom, and M. J. Fernstrom (1989). Vasopressin and
oxytocin receptors on plasma membranesfi-omrat mammary gland.
Demonstration of vasopressin receptors by stimulation of inositol
phosphate formation, and oxytocin receptors by binding of a specific
^25|-labeled oxytocin antagonist, d(CH2)5 ' [Try(Me)2. Thr^, Tyr-NH2T
OVT. Blochem. Cell Biol. 67:152-162.
SPSS, Inc. (1988). SPSS-X User's Guide. 3rd ed. SPSS Inc., Chicago, 1072 pp.
Stallone, J. N. and E. J. Braun (1988). Regulation of plasma antidiuretic
hormone in the dehydrated kangaroo rat (Dipodomys spectabilis M.).
Gen. Comp. Endocrln. 69:119-127.
Steiner, M. and M. I. Phillips (1988). Renal tubular vasopressin receptors
downregulated by dehydration. Amer. J. Physiol. 254:C404-C410.
Stevenson, J. L (1987). Models of the urinary concentrating mechanism. Kid.
Int 31:648-661.
Summy-Long, J., L M. Rosella-Dampman, G. L McLemore, and E. Koehler
(1990). Kappa opiate receptors inhibit release of oxytocin from the
magnocellular system during dehydration. Neuroendocrln. 51:376-384.
Tweedle, C. D. and G. 1. Hatton (1976). Ultrastructural comparisons of
neurons of supraoptic anc^ circularis nuclei in normal and dehydrated
rats. Brain Res. Bull. 1:103-121.
Vander Weele, D. A. , K. Schiffer, and D. L Esparza (1975).
Food-deprivation-induced polydipsia in two species of kangaroo rats.
Psych. Reports 36:645-646.
Vimtrup, B. J. and B. Schmidt-Nielsen (1952). The histology of the kidney of
kangaroo rats. Anat Rec. 114:515-528.
Wang, L C. and J. W. Hudson (1970). Some physiological aspects of
temperature regulation in the normothermic and torpid hispid pocket
mouse, Perognathus hispidus. Comp. Blochem. Physiol. 32:275-293.
Watt, R. M. (1970). Metabolic activity in single supraoptic neurones and its
relation to osmotic stimulation. Brain Res. 21:443-447.
51
Wood, A. E. (1935). Evolution and relationship of the heteromyid rodents with
new forms from the Tertiary of western North America. Ann. Carnegie
Mus. 24:73-262.
Wright, J. W., D. J. Gubernick, and T. J. Reynolds (1977). Intravascular,
carcass and gut fluid changes induced by food deprivation in xeric and
mesic adapted rodents. Comp. Blochem. Physiol. 58A:137-142.
Yagil, C. and C. D. Sladek (1990). Osmotic regulation of vasopressin and
oxytocin release Is rate sensitive in hypothalamoneurohypopysial
explants. Amer. J. Physiol. 258:R492-R500.
Yousef, M. K. and D. B. Dill (1971). Daily cycles of hibernation in the kangaroo
rat, Dipodomys merriami Cryoblol. 8:441-446.
52
APPENDIX
DESCRIPTIVE STATISTICS FOR PHYSIOLOGICAL PARAMETERS
53
Table A-1
Means ± standard errors for pituitary AVP (jig AVP/mg pituitary
weight). Column header * denotes differences between species in
the same treatment group. Superscript letters denote pairwise
comparisons of treatments within species; common letters show
pairs with no signiflcant differences between treatments.
Water
Stress
S*
Water
Control
W*
Baseline
Control
B*
C. hispidus
0.973 ±.075^
n=16
0.924 ±.065^
n=l7
1.017 ±.094^
n=18
D. ordii
0.434 ±.026^
n=20
0.442 ±.037^
n=19
0.088 ±.013
n=19
Species
54
^
Table A-2.
Means ± standard errors for plasma AVP (pg/ml). Column
header * denotes differences between species in the same
treatment group. Superscript letters denote pain/vise comparisons
of treatments within species; common letters signify no significant
differences between treatments.
Water
Control
Species
Baseline
Control
B*
W*
Water
Stress
S
C. hispidus
423 ± 3 5 ^
n=20
469 ± 4 9 ^
n=19
578 ± 4 2 ^
n=20
AsWii&ikmmm
1378 ± 1 0 7 ^
n=20
1010 ± 8 3 ^
n=18
818 ± 1 0 3 ^
n=19
55
1
1
Table A-3.
Means ± standard errors for urine osmolality (mOsm/kg). Column
header * denotes differences between species in the same
treatment group. Superscript letters denote pairwise comparisons
of treatments within species; common letters show pairs with no
significant differences between treatments.
Species
Water
Control
W
Baseline
Control
B*
i
Water
Stress
S
C. hispidus
1878 ±196^
n=20
1924 ±119^
n=20
3536 ±276
n=20
D. ordii
1158 ±126
n=20
2830 ±162^
n=20
3267 ±232^
n=18
56
Table A-4.
Means ± standard errors for plasma osmolality (mOsm/kg).
Column header * denotes differences between species in the
same treatment group. Superscript letters denote painvise
comparisons of treatments within species; common letters show
pairs with no significant differences between treatments.
'Ipiiii''
Water
Control
W*
Baseline
Control
B*
Water
Stress
S*
C. hispidus
320 ±0.9^
n=17
326 ±1.2^
n=18
310 ±2.9^
n=20
D, ordll
308 ± 1 . 1 ^
n=19
314 ± 1 . 1 ^
n=20
314 ± 1 . 8 ^
n=19
57
Table A-5.
Species
C hispidus
D. ordB
Means ± standard errors for hematocrit. Column header *
denotes differences between species in the same treatment
group. Superscript letters denote pairwise comparisons of
treatments within species; common letters show pairs with no
significant differences between treatments.
Water
Control
W
Baseline
Control
B
Water
Stress
S
45.2 ±0.7^
46.8 ± 0 . 7 ^
50.0 ±0.7^
n=18
n=17
n=17
44.5 ±0.7^
n=19
45.9 ± 0 . 7 ^
n=19
48.5 ±0.3^
58
n=16
1
Table A-6.
Species
Means ± standard errors for weight change (g). Column header
* denotes differences between species in the same treatment
group. Superscript letters denote pairwise comparisons of
treatments within species; common letters show pairs with no
significant differences between treatments.
Water
Control
W
Water
Stress
1
1
s*
I
C. hispidus
-f-0.44 ±0.41 s
n=20
-1.73 ±0.95^
n=20
D. ordii
-0.88 ±0.99
n=19
-7.33 ±0.78
n=19
59
Table A-7.
Species
C hispidus
D. ordii
Means ± standard errors for total pituitary AVP (jig AVP). Column
header * denotes differences between species in the same
treatment group. Superscript letters denote pain^/ise comparisons
of treatments within species; common letters show pairs with no
significant differences between treatments.
Water
Control
W*
Baseline
Control
B*
Water
j
Stress
S*
1
0.514 ±.073^
n=l6
0.488 ±.062^^
0.537 ±.095^
n=17
n=18
1.147 ±.030^
1.167 ±.040^
n=19
n=20
60
0.232 ±.013
n=19