Comparison of recovery from hemorrhage in birds and mammals

Camp. Biochem. Physiol. Vol. 87A, No. 3, pp. 533-542, 1987

0300-9629/87$3.00+ 0.00 0 1987Pergamon Journals Ltd

Printed in Great Britain

MINI

COMPARISON

REVIEW

OF RECOVERY FROM HEMORRHAGE IN BIRDS AND MAMMALS

S. L. SCHINDLER and R. P. GILDERSLEEVE* Poultry Science Department, Box 7608, North Carolina State University, Raleigh, NC 27695-7608, USA. Telephone (919) 737-2628 (Received 2 December

INTRODUCTION

The objective of this review is to compare the known physiological mechanisms involved in recovery from hypovolemia in mammals to those of birds. Most avian species recover plasma volume following hemorrhage more rapidly than mammals (Djojosugito et al., 1968; Kovach and Balint, 1969; Wyse and Nickerson, 1971; Schindler et al., 1987a). Moreover, it is generally accepted that most birds exhibit complete cellular recovery in response to blood loss in a shorter period of time than mammalian species. Chickens (Gulfus domesticus) recover total red blood cell numbers within 7 days after a 35% decrease in hematocrit due to hemorrhage (Natt and Herrick, 1955). Japanese quail (Coturnix coturnix juponica) recover total red blood cell numbers within 72 hours after loss of 30% of the estimated total blood volume (Gildersleeve et al., 1985a). The recovery period in mammals for less severe blood losses (5-15% total blood volume) is generally longer: 40 days in humans (Coleman et al., 1953); 14-17 days in rabbits (Smith and Weinberg, 1981); and 8-15 days in rats (Berlin and Lotz, 1951). The ability of birds to rapidly recover blood volume following a severe hemorrhage is reviewed herein by comparison to the well-known physiological compensations exhibited by hypovolemic mammals. Although the published literature on hemorrhagic shock and recovery in mammals is extensive, sparingly little literature on avian responses to hypovolemia is available. This review does not attempt to be a comprehensive account of the mammalian literature. However, it does represent an essentially comprehensive review of the avian literature. Blood pressure maintenance after hemorrhage

For many years researchers have characterized hemorrhagic hypotension by Wigger’s classical model of animal blood pressure maintained at a depressed level for a given period of time (Knott et al., 1969; Hess et al., 1983). The model has been criticized as not representing a naturally occurring hemorrhage (Knott et al., 1969). Models using single phlebotomies (Haddy et al., 1965; Gann and Pirkle, *To whom all correspondence

should be addressed. 533

1986)

1975), repeated small phlebotomies (Kovach et al., 1969) and single phlebotomies with partial replacement have been offered as alternative models (Knott et al., 1969). The use of different experimental models has made the study of this complex physiological event even more difficult, since results from various laboratories are very difficult to correlate. The study of hemorrhagic shock in mammals is further compounded by the sensitivity of most mammalian species to voluminous blood losses. The fact that mammals enter a phase of decompensation or irreversibility during hemorrhagic hypotension which cannot be ameliorated even by replacement of the lost blood is of great interest to researchers of mammalian hemorrhagic shock (Hess et al., 1983). The recovery of blood volume, although important for survival in mammals, is even more critical in avian species, since they depend upon blood volume replacement as their major blood pressure homeostatic mechanisms (Djojosugito et al., 1968). The recovery of avian species from hemorrhage has been studied to a much lesser degree than that in mammalian species. However, the lack of a standard model for the study of hemorrhage is also a difficulty when reviewing avian hemorrhagic recovery. Avian species, in general, are more resistant to blood loss than mammalian species. Flying and diving birds such as ducks exhibit the most resistance to hypovolemia (Kovach et al., 1969). Chickens maintained at a mean arterial pressure of 50 mmHg for three hours exhibited no phase of decompensation or irreversibility (Wyse and Nickerson, 1971). Chickens bled nearly to the point of cardiovascular collapse (3 hours at 50 mmHg) had a high survivability after reinfusion of the lost blood (Wyse and Nickerson, 1971). Acute blood loss results in a transient decrease in cardiac output due to decreased venous return, increased peripheral resistance and increased heart rate (Guyton, 1981). The immediate responses in mammals to acute hemorrhage is a reflex increase in peripheral resistance (Guyton, 1981), resulting in a peripheral to central shift in blood flow (Szlyk et al., 1984). The maintenance of blood pressure following hemorrhage in dogs is primarily due to cardiovascular reflexes causing a decrease in the total vascular space (Swan, 1965). Shrinking of larger

534

S. L. SCHINDLERand R. P. GILDER~LEEVE

blood vessels probably occurs in the venous system by reflex increases in venomotor tone in order to increase venous return and cardiac output (Swan, 1965; Szlyk et al., 1984). The response of the vascular system to hemorrhage is regulated by both neural and hormonal mechanisms. Reflex-increased venous tone appears to be mediated by alpha-adrenergic receptors (Szlyk et al., 1984). Alpha-ad~nergic mediated vaso~onst~ction has been reported to override myoge~c renal vasodilation in dogs subjected to severe hemorrhage (Kremser and Gewertz, 1985). Haddy et al. (1965) suggested that increased precapillary resistance was due to active vasoconstriction; specifically, by a baroreceptor-induced sympathico-adrenal discharge. This active vasoconstriction was not maintained in the limbs, intestines or kidney (Haddy et a/., 1965). Younes et al. (1985) observed that infusions of hypertonic saline solutions into venous blood supply caused increased arterial and pulse pressures that did not dissipate. However, when infused into the arterial blood supply, the effect of the hypertonic saline was not maintained. They concluded that for the response to occur, the hypertonic solution must pass by sinoreceptors located in the lung. These receptors apparently exert an important role in activating cardiovascular responses (Younes et al., 1985). Haddy et al. (1965) recognized that vasoconstriction in response to hemorrhage in dogs was due to circulating vasoconstrictors of both adrenal and nonadrenal origin. Sympathetic innervation is necessary for increased secretion of catecholamines in response to hemorrhage (Engleland et al., 1985). Catecholamines are strong vasoconstrictors (Guyton, 1981). Vasopressin and r~n/an~otensin are released in response to hemo~hage and are capable of causing intense vas~onst~ction (Goetz et al., 1984). Secretion of vasopressin in response to hemorrhage is mediated primarily through cardiac receptors (Goetz et al., 1984). Increased plasma renin activity is not due to increased stimulation of either sinoaortic or cardiac receptors (Goetz et al., 1984). Rocchini et al. (1985) attributed the initial vasoconstriction in dogs in response to hemorrhage to the actions of norepinephrine and vasopressin and maintenance of vasoconstriction to the activity of the renin/angiotensin system. Dogs fed chronic high salt diets prior to hemorrhage exhibited blunted renin/angiotensin responses to hemorrhage which resulted in decreased peripheral resistance, increased cardiac output and increased regional blood flow during later stages of shock (Rocchini et al., 1985). As in mammals, hemorrhage in avian species results in a drop in mean arterial pressure and increased heart rate (Kovach et al., 1969; Wyse and Nickerson, 1971; Zambraski and Schuler, 1980). Djojosugito et al. (1968) reported reduced blood flow to the lower leg muscle during hemorrhage in ducks (Anus plutyrhyncws), implying increased regional flow resistance to skeletal muscle. In contrast, Ploucha et al. (1981) reported that total peripheral resistance in the chicken either falls or remains the same during hemorrhage (Roucha and Fink, 1986) and that skeletal muscle flow resistance is unaffected. Homeostatic mechanisms in birds, as in mammals, work to maintain cardiac output when blood volume is suddenly

diminished (Kovach et al., 1969). The mechanisms by which certain avian species respond to blood loss are similar to those seen in mammalian species (Djojosugito et al., 1968) which use vasoconstrictive and blood volume replacement mechanisms. However, the degree to which avian species use these mechanisms is different from mammalian species (Djojosugito et a[., 1968). The vasocons~ctive m~hanisms in avian species, as in mammalian species, act to decrease vascular space and increase the pre- to postcapillary resistance ratio (Djojosugito et al., 1968). The pressor receptor zone of birds is located near the origin of the common carotid (Kovach et al., 1969). Unlike mammals, the tolerance of chickens to hemorrhage is not enhanced by prostaglandins which have vasoactive properties in mammals (Zambraski and Schuler, 1980). The ability of birds to withstand hemorrhage may be due to a less sensitive sympathetic-mediated vasoconstriction following acute blood loss. Kovach and Balint (1969) suggested that pigeons (Co/urn&r Ma) do not make full use of their ability to vasoconstrict in response to blood loss. inhibition of alpha-adrenergic receptors in birds failed to inhibit vasoconstriction stimulated by norepinephrine or epinephrine (Kovach and Balint, 1969). Further evidence supporting this view was provided by Ploucha et al. (1981). They reported that chickens did not exhibit the intense precapillary constriction seen in mammals. The relatively inefficient sympathetic adrenal response to hemorrhage by chickens may prevent deleterious effects due to inadequate tissue perfusion and aid in prevention of the phase of irreversibility seen in mammals (Ploucha et al., 1981; Ploucha and Fink, 1986). Hormonal vasoactive mechanisms have been suggested to play a role in recovery of birds following blood loss. Surgical adrenal~tomy resulted in progressive impairment of cardiovascular function in Pekin ducks (Butler and Wilson, 1985). Blood volume and hematocrit were not altered by adrenalectomy, while arterial blood pressure, stroke volume, osmolality, plasma concentrations of norepinephrine, epinephrine, Nat and Cl- were decreased (Butler and Wilson, 1985). Angiotensin II has been reported to increase blood pressure in ducks (Wilson and Butler, 1983a, b) and in chickens (Nishimura et al., 1982). Surgical adrenalectomy diminished the pressor repsonse to angiotensin II in ducks (Wilson and Butler, 1983a, b). The vasopressor actions of angiotensin II in ducks and chickens is probably due to stimulation of release of catecholamines from adrenal and extra-adrenal sources (Nishimura et al., 1982; Wilson and Butler, 1983a, b). In addition, Nishimura et al. (1982) reported a biphasic response to angiotensin II, that is, a pressor response preceded by a depressor response. The depressor response to angiotensin II may be the result of direct action of angiotensin II on smooth muscle and has been reported only in the chicken (Nishimura et al., 1982). Blood uolwne recovery after hemorrhage

Rapid recovery of blood volume following acute hemorrhage is recognized as an essential factor in m~ntai~ng the integrity of the cardiovascular system

Hemorrhage in birds during hemorrhagic shock in mammals (Adamson and Hillman, 1968; Gann et al., 1981). Increased cardiac output is a normal response to blood loss within the first 2 hours posthemorrhage (Szlyk ef al., 1984). The increased cardiac output is due to decreased blood viscosity and increased venous return (Szlyk et al., 1984). These postphlebotomy blood changes are due to an immediate hemodilution resulting from the influx of low-protein extravascular fluid (Adamson and Hillman, 1968). This influx of low-protein extravascular fluid is the first phase of blood volume replacement. Blood volume replacement following hemorrhage is characterized by two phases: a rapid, but still partial replacement during the first 15 minutes to 2 hours posthemorrhage (Pirkle and Gann, 1976) and a slower phase in which osmotic and volume recovery is completed. During the first phase of blood volume replacement, the influx of extravascular fluid into capillaries is due to decreased capillary hydrostatic pressure (Haddy et al., 1965). The factors causing this drop in capillary pressure are decreased aortic pressure, increased precapillary resistance and decreased right atria1 pressure (Haddy et al., 1965). The reason this first phase of blood volume recovery is only partial is due to decreased blood osmolality resulting from the influx of the low-protein extravascular fluid which offsets the decreased capillary hydrostatic pressure in mammals (Haddy et al., 1965). Swan (1965) reported that recovery of plasma volume during the first phase of recovery never exceeded 50% of the total blood volume removed and was usually much less than 50% in dogs. Movement of low-protein extravascular fluid across capillary membranes posthemorrhage also occurs in birds. The amount of absorption that takes place in the skeletal muscle of ducks is greater than that in cat skeletal muscle due to the presence of a larger capillary bed (Djojosugito et al., 1968). This could partially explain the greater resistance of birds to acute hemorrhage. However, if capillary hydrostatic pressure is offset by decreased osmotic pressure from hemorrhage-induced hemodilution in birds, as occurs in mammals, then this cannot be the only mechanism for blood volume restoration in birds. Birds must have a second stage of recovery just as in mammals. In two reports (Gildersleeve et al., 1985a; Schindler et al., 1987a), plasma protein levels were decreased in birds during the first six hours posthemorrhage. Plasma osmolality was also decreased immediately after hemorrhage in quail. In the study of Schindler et al. (1987a), the recovery of total erythrocyte volume and of plasma volume within one hour posthemorrhage resulted in complete recovery of total blood volume within one hour posthemorrhage. This is less than 5% of the average time necessary for recovery in most mammalian species. It is, therefore, unlikely that protein played a significant role in the rapid postphlebotomy recovery of plasma volume in quail. The ability of pigeons to survive slow blood losses equivalent to their initial blood volume (Kovach and Balint, 1969) suggests that blood volume replacement is of primary importance to the survival of birds during hemorrhage. Djojosugito et al. (1968) re-

535

ported that ducks showed almost complete restoration of blood volume within 20 to 25 minutes after a 13 to 15 percent blood loss. After 3 hours at a depressed blood pressure of 50 mmHg, plasma volume in chickens was not significantly reduced. An average fluid mobilization equivalent to 60 percent of the initial plasma volume was mobilized to expand the plasma volume during hemorrhage (Wyse and Nickerson, 1971). The two main factors in recovery of blood volume are a net transfer of fluid from the extravascular fluid and a decreased fluid loss in the kidneys (Kovach and Balint, 1969). The ability of birds to replace blood volume very rapidly may be due to greater capillary surface area in skeletal muscle and reflex lowering of mean capillary pressure (Djojosugito et al., 1968). Djojosugito et al. (1968) reported a rapid, steady decrease in the lower leg muscle volume of hemorrhaged ducks reflecting a loss of fluid from the skeletal muscle. This fluid loss was reduced by 75% when vasomotor blockade with xylocaine was imposed. This indicates that neural control for fluid shift can occur in certain birds (Djojosugito et al., 1968). The second phase of blood volume recovery occurs more slowly in mammals. Most species studied completely recover blood volume within 24 hours posthemorrhage (Knott et al., 1969; Gann et al., 1981). However, Adamson and Hillman (1968) reported that humans in which 15-20% of the total blood volume was removed required at least 72 hours for complete blood volume recovery. One of the essential factors necessary for complete blood volume replacement in mammals during the second phase is recovery of colloid osmotic pressure by rapid return of protein to the plasma from the lymphatic system (Cope and Litwin, 1962; Gann et al., 1981). The increased osmotic pressure due to the recovery of plasma protein potentiates augmentation of plasma volume with extravascular fluid (Cope and Litwin, 1962). After total blood volume is restored, Japanese quail recover plasma protein within 72 hours postphlebotomy (Gildersleeve et al., 1985a). Avian species do not have a well developed lymphatic system (Sturkie, 1976), so rapid recovery of colloid osmotic pressure is unlikely to be the osmotically active agent in the second stage of blood volume recovery. Although most researchers generally agree that plasma protein replacement postphlebotomy is the main osmotic driving factor during the second phase of blood volume recovery, other osmotically active agents have been suggested to play important roles in increasing plasma osmolality postphlebotomy in both birds and mammals. Hyperglycemia has been reported as the main factor in postthemorrhage hyperosmolality of plasma (Knott et af., 1969; Jarhult, 1973). The principle cause of the hyperglycemia is increased glycogenolysis by the liver in response to increased circulating levels of catecholamines (Knott et al., 1969; Jarhult, 1973). Pirkle and Gann (1976) suggested that glucose plays an important role in plasma hyperosmolality during hemorrhagic shocks, but is of minor importance in rapid, moderate hemorrhages. Hyperglycemia following massive hemorrhages in dogs may play an important role in potentiating plasma volume recovery (Pirkle and Gann, 1976). It appears that

536

S. L. SCHINDLER and

hyperglycemia also plays a major role in Japanese quail in the recovery of plasma volume (Gildersleeve et al., 1985a; Schindler et al., 1987a). Hemorrhage is known to increase norepinephrine, glucocorticoid and glucagon secretion and liver glycogenolysis which results in blood glucose level increases (Chaudry and Baue, 1982). Early support for a role of glucocorticoids (cortisol) in mediating increased osmolality postphIebotomy was reported by Marks et al. (1965). They reported that surgically adrenalectomized dogs were capable of maintaining blood pressure and increasing plasma volume following small hemorrhage only with administration of glucocorticoids. Gann and Pirkle (1975) suggested that inhibition of amino acid uptake of cells other than liver cells by increased circulating levels of cortisol released during hemorrhage may play an active role in postphlebotomy hy~rosmolality. Increased plasma concentrations of free amino acids may be important osmotic agents (Gann and Pirkle, 1975). However, glucose, Na+ and K+ are not mobilized by increased levels of plasma cortisol posthemorrhage (Pirkle and Gann, 1976). Another important osmotically active substance in plasma is Na+. Knott et al. (1969) reported that plasma Na+ concentrations remained constant with increasing plasma volumes postphlebotomy, indicating increased plasma Na+ concurrent with increased plasma volume. They suggested that this was due to either increased renal absorption, decreased renal clearance or release of endogenous sources of Na+ (Knott et OZ., 1969). Schindler et al. (1987a) reported immediate but transient rises in plasma Na+ after hemorrhage in quail which were regulated rapidly down to normal ranges. In addition, no change in osmolality and plasma Na+ with concurrent hemodilution reflected rapid regulation of these parameters during hemorrhagic hypotension in chickens (Ploucha et al., 1981; Nouwen et al., 1984). Aldosterone is the major natrieuretic hormone in mammals. Plasma aldosterone has also been reported to increase following hemorrhage (Ganong and Mulrow, 1962). The increased release of aldosterone by the adrenal occurs in response to ACTH release by the anterior pituitary during hemorrhage (Ganong and Mulrow, 1962). In addition, release of aldosterone during hemorrhage may be activated in hypophysectomized dogs, indicating an alternative mechanism of aldosterone release (Ganong and Mulrow, 1962). This alternative mechanism is probably the renin-angiotensin system (Ganong and Mulrow, 1962). Recent evidence of an atria1 natriuretic factor (ANF) produced by cardiac atrial muscle cells and which inhibits renin and aldosterone secretion (de Bold, 1985) confirms the presence of multiple factors regulating kidney water and electrolyte secretion. However, a role for ANF during hemorrhagic shock has not been elucidated. Results from salt-loading and water deprivation experiments and evidence of positive correlations between plasma Nat and plasma prolactin suggest that prolactin and arginine vasotocin play important osmoregul~tory roles in domestic fowl (Nouwen et al., 1984). The main target organ of prolactin may be the intestine, although this does not rule out a

R. P. GILDERSLEEVE possible synergistic action of prolactin with arginine vasotocin in the kidney (Nouwen et al., 1984). Increased levels of another neurohypophyseal peptide, mesotocin, during hypovolemia suggests that it plays a role in volume regulation in the chicken (Nouwen et al., 1984). As in mammals, hemorrhage in chickens stimulates aldosterone and glucocorticoids (Radke et al., 1985). Gildersleeve (unpublished~ found increased plasma corticosterone levels in response to hemorrhage in Japanese quail. This adrenal response by birds is partially due to ACTH stimulation, although the aldosterone response may be due in part to other factors (Radke et al., 1985). It is possible that part of the aldosterone response to hemorrhage is due to increased stimulation of the renin-angiotensin system (Radke et al., 1985). In Japanese quail this is supported by evidence that injections of angiotensin II induce increases in plasma aldosterone, arginine vasotocin, and corticosterone (Kobayashi and Takei, 1982). In addition to kidney and skeletal muscle capillary fluid shift responses to hypovolemia, Japanese quail have been reported to drink water following hemorrhage or administration of hypertonic saline solution (Kobayashi and Takei, 1982). Schindler et al. (1987b) measured feed and water consumption in response to a hemorrhage via phlebotomy of 30% of the calculated total blood volume with and without replacement of blood volume with isotonic physiological saline in juvenile male Japanese quail. Feed consumption was not affected by phlebotomy or phlebotomy with saline replacement. Water consumption of the hemorrhaged quail was higher than nonhemorrhaged controls during the 24 hr postphlebotomy period. This supports previous reports of vigorous drinking within 30 minutes posthemorrhage by Japanese quail (Kobayashi and Takei, 1982) and suggests that gut absorption of water may be another mechanism by which quail replenish plasma volume postphlebotomy. Replacement therapy with plasma substitutes following massive blood loss has been studied in mammals and birds (Schindler et ui., 1987b). Expansion of plasma volume following blood losses may lead to an increased survival following hemorrhage. Lawson and Rehm (1943) reported no difference between plasma and serum as volume expanders after a single rapid hemorrhage. Swingle et al. (1944) reported that intermittent replacement with a gelatin solution was more effective than a single massive replacement in preventing death and that saline replacement was less effective than gelatin. Cope and Litwin (1962) suggested that plasma expansion with solutions such as physiological saline, which were freely permeable to capiliary membranes, are ineffective in restoring plasma volume post-hemorrhage. Crystalloid plasma expanders also have been utilized in treatment of hemorrhagic shock in dogs. Dextran 40, Dextran 70 and lactated Ringer’s solution have been reported to correct the effects of hemorrhagic shock in dogs comparable to the effect of plasma expansion by colloidal solutions (Gross et al., 1984). Plasma expansion utilizing 10% dextrose solution tended to have a decreasing cardiac output 15 to 30 minutes posttreatment (Gross er al., 1984).

Hemorrhage in birds Hematopoiesis after hemorrhage or hypoxia

Cellular recovery from anemias induced by phlebotomy or hemolysis has been well studied in mammals. Schalam (1965) in his textbook Veterinary Hematology has presented a comprehensive review of hemopoiesis in domestic mammals. Erythropoiesis occurs primarily in bone marrow, with the amount of bone marrow actively involved in erythropoiesis decreasing as young animals mature until only the medullary bones maintain erythropoietic capability. The loss of erythropoietic capability by bone marrow is characterized by a morphological change from red marrow (erythropoietic) to fatty yellow marrow. In times of great demand upon the bone marrow for erythrocyte production such as acute hemorrhage or hemolysis, yellow marrow is capable of recrudescing into erythropoietically active red marrow within 48 hours after the hemorrhagic or hemolytic event. In addition to the bone marrow, the spleen also retains its capacity for hemopoiesis and may play an active part in recovery from blood loss in addition to its role as a red blood cell reservoir (Schalam, 1965; Lord and Murphy, 1973). The spleen may be the major site of erythropoiesis in response to hypoxia in some strains of mice (Shadduck et al., 1969). The attempts to understand the sites of erythropoiesis in avian species were similar to those in mammals (Lucus and Jamroz, 1961). Additionally, the conclusions for these studies indicated that birds produced red blood cells in sites similar to those in mammals. However, this assumption may not be tenable because mature avian erythrocytes retain their nucleus (Lucas and Jamroz, 1961). Erythropoiesis of sexually mature birds is confined primarily to the bone marrow (Glick and Rosse, 1976). In addition, erythropoiesis in the peripheral circulation of chickens by a “cloning” mechanism of young mature erythrocytes has been reported (Smith and Engelbert, 1969). Apparently, no literature is available on the erythropoietic ability of the spleen, liver or kidney of any mature avian species. Increased erythrocyte production, which can be recognized by an increased number of reticulocytes released into circulation, has been observed in humans and all domestic and laboratory animals except the horse in response to anemia (Giblett et al., 1956; Smith and Agar, 1975). The reticulocyte response is initiated in response to loss of red blood cells or exposure to a hypoxic environment. However, in some strains of mice bred for low erythropoietic response to hypoxia, normal erythropoietic responses are seen when anemia is induced by phlebotomy. Apparently, two separate mechanisms exist in this animal for hemopoietic recovery (Lord and Murphy, 1973). In human bone marow, a large reserve supply of reticulocytes can be released into peripheral circulation in addition to an increased erythropoiesis in response to anemia (Giblett et al., 1956). Shadduck et al. (1969) reported that certain strains of mice responded to hypoxia with increased splenic erythropoiesis, while bone marrow erythropoiesis remained at basal maintenance levels. The ability of the erythropoietic organs to respond to anemia or hypoxia is dependent on adequate dietary iron supply (Hillman and Henderson, 1969). When a large demand for erythropoiesis is placed

537

on the bone marrow, immature reticulocytes and even normoblasts, the progenitor cell of the reticulocyte, may be released into circulation (Giblett et al., 1956; Hillman, 1969). These immature or “stress” reticulocytes are released following hemorrhage in the rabbit (Borsook et al., 1962). Borsook et al. (1962) suggested that these reticulocytes arise directly from polychromatic erythroblasts by skipping the mitotic subdivision stage of development. This results in the release of reticulocytes that are larger and contain more hemoglobin than normal. Stress reticulocytes also synthesize protein at a higher rate than normal reticulocytes (Smith and Weinberg, 1981). In dog bone marrow, the spleen may play an inhibitory role by preventing the release of stress reticulocytes (Lorber, 1961). Circulating stress reticulocytes mature into erythrocytes in 3 to 4 days in the rat and 1 to 2 days (Smith and Weinberg, 1981) in the rabbit. These “stress” erythrocytes occur in adequate numbers to replace the lost red blood cells, but have shorter than average life spans in circulation in the rat (Berlin and Lotz, 1951; Stohlman, 1961b) and the rabbit (Neuberger and Niven, 1951). Neuberger and Niven (1951) suggested that the shortened life span of stress erythrocytes is due to skipping stages in the erythroid cell maturational process and Stohlman (1961b) suggested that the macrocytic character of stress erythrocytes plays a role in their decreased life span in circulation. The avian response to hemolytic or hemorrhagic anemias includes increased erythropoiesis by the bone marrow which is characterized by increased numbers of circulating reticulocytes (Natt and Herrick, 1955; Rosse and Waldmann, 1966; Stino and Washburn, 1970; Jones et al., 1978; Gildersleeve et al., 1985a,b; Clark et al., 1987a). Clark et al. (1987b) reported reduced circulating erythroblast numbers postphlebotomy in Japanese quail. Unlike mammals, the spleen of the chicken does not act as a reservoir of mature red blood cells than can be released into circulation in response to anemia (Sturkie, 1943). No evidence exists that any organ other than the bone marrow can respond to anemia with erythropoiesis. The reticulocytes released in response to acute hemorrhage mature into erythrocytes within 1 to 3 days in Japanese quail (Gildersleeve et al., 1985a). This recovery time is comparable to reticulocyte maturation in mammals. It is not known if the erythrocytes produced in response to phlebotomy in birds have a shortened life span. However, Gildersleeve et al, (1985a, b) reported that mean cell volume, mean cellular hemoglobin and mean cellular hemoglobin concentration were elevated, while hemoglobin was depressed during the first 6 hours postphlebotomy in Japanese quail. By 72 hours postphlebotomy, all hematological values except mean cellular hemoconcentration had returned to nonglobin phlebotomized levels. Hematological responses to hemorrhage by phlebotomy with and without replacement of blood volume with isotonic saline were determined in Japanese quail by Schindler et al. (1987b). Recovery of total peripheral erythrocyte numbers within 72 hours postphlebotomy occurred in both treatment groups. Saline replacement of blood volume following hemorrhage increased the total numbers and differential

538

S. L.

SCHINDLERand

percentages of circulating reticulocytes at 72 hours postphlebotomy above the reticulocyte values of phlebotomized quail receiving no saline in both adult and juvenile Japanese quail. The last 80 years of research on erythropoietin in mammals has been reviewed recently by Jelkmann (1986) and it is generally agreed that the stimulation of erythropoiesis is primarily under hormonal control in mammals. Jacobson et al. (1957) suggested that synthesis of erythropoietin is related to changes in the relation of oxygen supply to oxygen demand and reported great increases in plasma erythropoietin by 10 to 12 hours after an acute hemorrhage. Fried et al. (1956) reported that plasma from hemorrhaged rats previously subjected to hypophysectomy, thyroidectomy, splenectomy, adrenalectomy or gonadectomy was still capable of producing increased erythropoiesis in intact rats comparable to the response produced by plasma from intact anemic rats. In addition, removal of the adrenals, pancreatic, stomach, intestines or approximately 90% of the liver did not affect the increase in plasma erythropoietin in response to a massive hemorrhage (Jacobson et al., 1957). However, bilateral nephrectomized rats and rabbits were not capable of responding to hemorrhage with increased levels of plasma erythropoietin as measured by stimulation of erythropoiesis in intact animals by injection of plasma from nephrectomized animals. These results support the hypothesis that erythropoietin is produced by the kidney. Based on data from hypoxic rats, Katz et al. (1968) offered an alternative hypothesis; specifically, the kidney is stimulated to secrete an activator which converts a plasma substrate of liver origin into the active form of erythropoietin during the initial phase of hypoxia. In vitro studies support a model of a renal source of erythropoietin synthesis (Jelkmann, 1986). Synthesis of erythropoietin may be stimulated by renal hypoxia which activates oxygen-sensitive cells on the venous side of the renal capillary network to stimulate synthesis of erythropoietin. However, other authors suggest that the synthesis of erythropoietin in response to decreased blood-oxygen tension is not the only means of stimulation of erythropoiesis (Stohlman, 1959; Stohlman, 1961a). Stohlman (1959) offered an alternative hypothesis; synthesis of an inhibitor substance by red blood cells which is sequestered in the cell as it ages and, upon normal destruction of the mature cell, is released and acts directly on the bone marrow. Removal of a large number of mature cells by bleeding would decrease the amount of inhibition upon the bone marrow and allow increased erythropoiesis. Attempts to verify this hypothesis have been unsuccessful (Stohlman, 1959). Shadduck et nl. (1968) reported that a strain of mice (CAF,) responded to hemorrhage with a normal increase in plasma erythropoietin, but responded to hypoxia with only a minimal increase in plasma erythropoietin. This suggests the existence of two separate mechanisms of erythropoietin stimulation. Rats maintained in the head down suspension showed a decreased response of erythropoietic tissue to erythropoietin which was attributed to food and water deprivation resulting from the restraint of the animals (Dunn et al. 1986). Splenic erythropoiesis was affected more than the erythropoietic activity of

R. P.

GILDERSLEEVE

the bone marrow in those animals. This finding also suggests different sensitivities of erythropoietic tissues. Rich et al. (1982) reported that macrophages in mice continually synthesize and release erythropoietin. They suggested that Kupffer cells in the liver are the main extrarenal source. In addition, macrophages could play a direct cell to cell role in the regulation of erythropoiesis in the bone marrow (Rich et al., 1982). Rosse and Waldmann (1966) first reported the presence of an erythropoietic factor in serum of anemic chickens and hypoxic quail. The “avian erythropoietin” differed from mammalian erythropoietin in that it did not stimulate erythropoiesis in polycythemic mice and was not bound by an antibody to human erythropoietin. In addition, human erythropoietin did not stimulate erythropoiesis in Japanese quail (Rosse and Waldmann, 1966). The erythropoietic factor in serum of anemic chickens was partially purified by Samarut (1978). After erythropoietin is secreted, the time required for complete recovery of red blood cell number varies between species and is dependent upon the severity of the anemia. The average time for recovery from a blood volume loss of 5-15% is 40 days in humans (Coleman et al., 1953), 14 to 17 days in rabbits with phenylhydrazine-induced hemolysis (Smith and Weinberg, 1981) and 8 to 15 days in hemorrhaged rats (Berlin and Lotz, 1951). Recovery of red blood cell numbers is generally more rapid in avian species than in mammals. Stino and Washburn (1970) reported that increased reticulocytosis was recognizable by 3 days postinjection of phenylhydrazine in chickens and by 24 hours postinjection in quail (Clark et al., 1987a). Chickens also recover total red blood cell numbers and hematocrit after a 35% decrease in hematocrit within 7 days postphlebotomy (Natt and Herrick, 1955). Japanese quail have been reported to respond to phlebotomies of 30% of the estimated total blood volume with increased reticulocytes within 24 hours postphlebotomy and recovery of total red blood cell and hematocrit to 0 hour (nonnumbers phlebotomized) levels by 72 hours postphlebotomy (Gildersleeve et al., 1985a). The red blood cells remaining in circulation after hemorrhage, hemolysis or during exposure to a hypoxic environment also respond to the change in oxygen tension of blood. Erythrocytes in mammals respond to hypoxia with increased synthesis of 2,3-diphosphoglycerate which acts to decrease the affinity of hemoglobin for oxygen resulting in a shift of the oxy-hemoglobin dissociation curve to the right (Oski and Gottlieb, 1971). This results in increased oxygen delivery to body tissues. The increased oxygen delivery due to increased synthesis of 2,3-diphosphoglycerate, however, is not sufficient to compensate for decreased total oxygen carrying capacity due to massive hemorrhage or hemolysis (Torrence et al., 1970; Naylor et al., 1972; Holter and Refsum, 1985). Metabolic responses of mature erythrocytes to hypoxia in avian species also differs from mammals. Vandecasserie et al. (1971) reported that avian and turtle erythrocytes contain inositol-hexaphosphate

Hemorrhage in birds instead of 2,3-diphosphoglycerate and that inositolhexaphosphate has a regulatory effect on the oxygen affinity of hemoglobin in the avian and reptilian erythrocyte. The main organic phosphate in avian erythrocytes that earlier researchers identified as inositol-hexaphosphate has since been identified as inositol-pentaphosphate (Jones et al., 1978; Isaacks and Harkness, 1980). Jones et al. (1978) reported rapid postphlebotomy rises in mature chicken erythrocyte inositol-pentaphosphate which would act as 2,3-diphosphoglycerate in mammals to decrease the affinity of hemoglobin for oxygen and allow more oxygen to be delivered to tissues. In contrast, Isaacs et al. (1983) reported that levels of inositolin chicken erythrocytes postpentaphosphate hemorrhage do not increase above normal levels found in mature chicken erythrocytes. In addition, avian red blood cells are unable to catabolize inositolpentaphosphate once it is formed (Isaacks et af., 1983). From this study, Isaacks et al. (1983) suggested that inositol-pentaphosphate levels were not altered as readily as 2,3-diphosphoglycerate was in mammals in response to anemia or hypoxia. The leukocyte response to hemorrhage includes an increased number of neutrophils in circulation (Guyton, 1981). Knott et al. (1969) reported no changes in leukocyte values and no change in white cell differentials in dogs in response to acute hemorrhage. The maintenance of leukocyte values within normal ranges concurrent with dilution due to plasma replacement reflects increased synthesis of white blood cells in response to hemorrhage (Knott et al., 1969). Synthesis of mammalian polymorphonuclear white cells, distinguishable by the presence of granules in their cytoplasm, occurs primarily in the bone marrow. Megakaryocytes, the precursor of thrombocytes, are also found only in the bone marrow, while lymphocytes and plasma cells are synthesized in lymphoid tissues throughout the body but not in the bone marrow (Schalam, 1965; Guyton, 1981). In times of high cellular demand from the bone marrow, the spleen is also capable of producing granulocytes and megakaryocytes (Schalam, 1965). Early reports that the bone marrow has a high amount of lymphoid tissue (Lucas and Jamroz, 1961) have been contradicted by other reports of minimal amounts of bone marrow lymphoid tissue (Glick and Rosse, 1976) in the chicken. Glick and Rosse (1976) reported that granulopoiesis in the chicken occurs in the bone marrow as it does in mammals, but unlike that of mammals, chicken bone marrow does not contain large reserves of mature granulocytes. The role of the spleen in granulopoiesis appears to be only a reserve for lymphoid tissue (Lucas and Jamroz, 1961). Megakaryocytes have not been observed in the bone marrow of any avian species, although thrombocytes are present in peripheral circulation (Schalam, 1965) and are reported to be the primary circulating phagocyte in chickens (Chang and Hamilton, 1979). To the author’s knowledge, the only studies reporting avian leukocyte changes postphlebotomy are those of Gildersleeve et al. (1985b), Schindler et al. (1986b), and Clark et al. (1987b). Gildersleeve et al. (1985b) reported lymphocyte recovery within 24 hours in adult quail and within 48 hours in juvenile C.B.P. 87,3&-B

539

quail. Heterophil numbers were unaffected at 6 hours postphlebotomy indicating a rapid recovery, but decreased well below nonphlebotomized levels thereafter in adult males and juvenile females. No consistent postphlebotomy effects on monocytes, eosinophils, or basophils were seen. Generally, total white blood cell recovery was more rapid than red cell recovery. Schindler et al. (1987b) and Clark et al. (1987b) observed heterophilia at 6 hr posthemorrhage in hemorrhaged birds. This agrees with previous reports reviewed by Siegel (1971) in which heterophilia occurred in response to stress or ACTH administration. Gildersleeve et al. (unpublished) found elevated plasma corticosterone levels in quail after hemorrhage and Radke et al. (1985) reported no difference in the corticosterone increase in hemorrhaged or restrained but not hemorrhaged chickens. Schindler et al. (1987b) and Clark et al. (1987b) also observed the recovery of total leukocyte numbers by 72 hr postphlebotomy which agrees with previous reports of leukocyte recovery postphlebotomy (Gildersleeve et al., 1985a). The rapid recovery of total leukocyte numbers may be due to release of sequestered leukocytes by the bone marrow. Lucas and Jamroz (1961) reported large amounts of lymphoid tissue in avian bone marrow. However, Glick and Rosse (1976) did not find large amounts of lymphoid tissue in chicken bone marrow and reported that chicken bone marrow did not contain the large reserves of mature granulocytes found in mammalian bone marrow. CONCLUDING

REMARKS

Although reports in avian species characterizing cellular responses to hemorrhagic and hemolytic anemias are available, a paucity of information on the regulation and control of erythropoiesis in birds is available. Whether tissue hypoxia or another stimulus such as a rapid drop in mean arterial pressure or loss of a plasma constituent is the main stimulus of erythropoiesis after acute blood loss has not been elucidated. In addition, the interaction among certain events following hemorrhage has not been determined in either mammalian or avian species. That is, whether blood volume recovery is necessary prior to cellular recovery or if recovery of a plasma constituent such as Na+ or protein affects initiation of cellular recovery has yet to be determined in either mammalian or avian species. The rapid recovery of blood volume in birds, which lack a well-developed lymphatic system, suggests that factors other than rapid recovery of plasma protein are involved in blood volume recovery. What factors are responsible for the rapid replacement of blood volume in birds have not been elucidated. Before an accurate comparison of mammalian and avian blood volume recovery mechanisms can be made, the factors regulating blood volume replacement must be better understood. The mammalian system, which seems to be the less effective, has been extensively studied. The avian system of blood volume recovery has received little attention, yet has been shown to be a more effective mechanism for blood volume replacement. Therefore, additional studies of blood

540

S. L. SCHINDLER and

volume recovery should be directed toward elucidating the regulatory mechanisms of birds to better understand their ability to survive massive blood loss. An excellent comparative study of the hemodynamics of hemorrhage in the conscious rat and chicken has been published recently (Ploucha and Fink, 1986). Acknowledgements-The authors are pleased to acknowledge the assistance of T. E. Bryan and C. Smith in the preparation of the manuscript. This work was supported in part by NIEHS contract No l-ES-2-5019. Paper No. 10791 of the Journal Series of the North Carolina Agricultural Research Service, Raleigh, NC 27695-7601. The use of trade names in this publication does not imply endorsement by the North Carolina Agricultural Service of the products names, nor criticism of similar ones not mentioned.

REFERENCES Adamson J. and Hillman R. S. (1968) Blood volume and plasma protein replacement following acute blood loss in normal man. J. Am. Med. Assoc. 205, 6099612. Berlin N. I. and Lotz C. (1951) Life span of the red blood cell of the rat following acute hemorrhage. Proc. Sot. exp. Biol. Med. 78, 788-790.

Borsook H., Lingrel J. B., Scare J. L. and Millette R. L. (1962) Synthesis of hemoglobin in relation to the maturation of erythroid cells. Nurure l%, 347-350. Butler D. G. and Wilson J. X. (1985) Cardiovascular function in adrenalectomized Pekin ducks (Anus plufyrhynchos). Comp. Biochem. Physiol. MA, 353-358.

Chang C. F. and Hamilton P. B. (1979) The thrombocyte as the primary circulating phagocyte in chickens. J. Reticuloendothelial Sot. 25, 585-590.

Chaudry I. H. and Bave A. E. (1982) Pathophysiology of Shock, Anoxiu and Ischemia, pp. 203-219. Williams and Williams, Baltimore, MD. Clark M. W., Gildersleeve R. P., Thaxton J. P., Parkhurst C. R. and McRee D. I. (1987a) Hematological effects of ethyl methanesulfonate, paraquat and phenylhydrazine in Japanese quail. Comp. Biochem. Physiol., submitted. Clark M. W., Gildersleeve R. P., Thaxton J. P., Parkhurst C. R. and McRee D. I. (1987b) Leukocyte numbers in hemorrhaged Japanese quail after microwave irradiation in ova. Comp. Biochem. Physiol. in press. Coleman D. H., Stevens Jr A. R., Dodge H. T. and Finch C. A. (1953) Rate of blood regeneration after blood loss. Arch. Intern. Med. 92, 341-349. Cope 0. and Litwin S. B. (1962) Contribution

of the lymphatic system to the replenishment of the plasma volume following a hemorrhage. Ann. Surg. 156,655-667. de Bold A. J. (1985) Atria1 natriuretic factor: A hormone produced by the heart. Science (Wash. DC) 230,767-770. Djojosugito A. M., Folkow B. and Kovach A. G. B. (1968) The mechanisms behind the rapid blood volume restoration after hemorrhage in birds. Acta Physiol. Stand. 74, 114-122. Dunn C. D. R., Johnson P. C. and Lange R. D. (1986) Regulation of hematopoiesis in rats exposed to antiorthostatic hypokinetic/hypodynamia. II. Mechanisms of the “anemia”. Aviat. Space Environ. Med. 57, 36-44. Engleland W. C., Lilly M. P. and Gann D. S. (1985) Symphathetic adrenal denervation decreases adrenal blood flow without altering the cortisol response to hemorrhage. Endocrinology 117, lOO&lOlO. Fried W., Plzak L., Jacobson L. 0. and Goldwasser E. (1956) Erythropoiesis. II. Assay of erythropoietin in hypophysectomized rats. Proc. Sot. exp. Biol. Med. 92, 203-207.

Gann D. S., Carbon D. E., Bymes G. J., Pirkle Jr. J. C. and Allen-Rowland

C. F. (1981) Impaired restitution

of

R. P. GILDERSLEEVE blood volume after large hemorrhage.

J.

Trauma 21,

598603.

Gann D. S. and Pirkle Jr J. C. (1975) Role of cortisol in the restitution of blood volume after hemorrhage. Am. J. Surg. 130, 565569.

Ganong W. F. and Mulrow P. J. (1962) Role of the kidney in adrenalcortical response to hemorrhage in hypophysectomized dogs. Endocrinology 70, 182-188. Giblett E. R., Coleman D. H., Pirzio-Birold G., Donohue D. M., Motulsky A. G. and Fionch C. A. (1956) Erythrokinetics: Quantitative measurements of red cell production and destruction in normal subiects and patients with anemia. Blood 11, 291-309. Gildersleeve R. P.. Galvin M. J.. Thaxton J. P. and McRee D. I. (1985a) Hematological response of Japanese quail to acute hemorrhagic stress. Comp. Biochem. Physiol. 81A, 403-409.

Gildersleeve R. P., Phelps P. V., Thaxton J. P. and McRee D. I. (1985b) Effect of phlebotomy on reticulocyte numbers in Japanese quail. Poultry Sci. 64, 199&1995. Glick B. and Rosse C. (1976) Cellular composition of the bone marrow in the chicken. I. Identification of cells. Anat. Rec. 185, 2355246. Goetz K., Wang B. C. and Sundet W. D. (1984) Comparative effects of cardiac receptors and sinoartic baroreceptors on elevations of plasma vasopressin and renin activity elicited by hemorrhage. J. Physiol. (Paris) 79, 440-445.

Gross D. R., Dodd K. T., Welch D. W. and Fife W. P. (1984) Hemodynamic effects of 10% dextrose and Dextran 70 on hemorrhagic shock during exposure to hyperbaric air and hyperbaric hyperoxia. Aviat. Space Environ. Med. 55, 111811128. -Guvton A. C. (1981) Textbook of Medical Phvsioloev. DD. g&64,367-368,396,4355445. W. B. Saunders Co&ah;, Philadelphia, PA. Haddy F. J., Scott J. B. and Molnar J. I. (1965) Mechanism of volume replacement and vascular constriction following hemorrhage. Am. J. Physiol. 208, 1699181. Hess M. L., Warner M. and Okabe E. (1983) Handbook of Shock and Trauma Vol. 1: Basic Science (Edited by B. M. Altura et al.), pp. 3933412. Raven Press, New York. Hillman R. S. and Henderson P. A. (1969) Control of marrow production by the level of iron supply. J. Clin. Invest. 48, 454460.

Hillman R. S. (1969) Characteristics of marrow production and reticulocyte maturation in normal man in response to anemia. J. Clin. Invest. 48, 4433453. Holter P. H. and Refsum H. E. (1985) Erythrocyte 2,3-diphosphoglycerate and erythropoietic activity in rabbits with severe bleeding anemia superimposed on the early post-natal fall in hemoglobin. Acta Phyisol. Scund. 124, 543-547.

Isaacks R. E. and Harkness D. R. (1980) Erythrocyte organic phosphates hemoglobin function in birds, reptiles and fishes. Am. Zool. 20, 115-129. Isaacks R., Kim C., Liu H. L., Goldman P., Johnson Jr A. and Harkness D. (1983) Studies on avian erythrocyte metabolism. XIII. Changing organic phosphate composition in age-dependent density populations of chicken krythrocytes-Poultry Sci. 62, 1639-i646. Jacobson L. 0.. Goldwasser E.. Fried W. and Plzak L. (1957) Role of the kidney in erythropoiesis. Nature 179, 633-634.

Jarhult J. (1973) Osmotic fluid transfer from tissue to blood during hemorrhagic hypotension. Acta Physiol. Stand. 89, 213-226.

Jelkmann W. (1986) Erythropoietin research, 80 years after the initial studies by Camot and Deflandre. Resp. Physiol. 63, 257-266.

Jones S. R., Smith J. E. and Board P. B. (1978) Changes in erythrocyte metabolism following acute blood loss in chickens. Poultry Sci. 57, 1667-1674.

Hemorrhag : in birds Katz R., Cooper G, W., Gordonn A. S. and Zanjani E. D. (1968) Studies on the site of production of erythropoietin. Ann. N.Y. Acad. Sci. 148, 120-127. Knott D. H., Beard J. D. and Overman R. R. (1969) The response of the dog to repeated acute hemorrhages. Am. Surg. 35, 284-29 1. Kobayashi H. and Takei Y. (1982) Mechanisms for induction of drinking with special reference to angiotensin II. Cornp. Biochem. Physiol. 71A, 485-494. Kovach A. G. B. and Balint T. (1969) Comparative study on haemodilution after hemorrhage in the pigeon and the rat, Acra Phvsiol. Acad. Sci. Huna. 35. 231-243. Kovach A. G. i., Szasz E. and Pilmaier i. (1969) Mortality of various avian and mammalian species following blood loss. Acta Physiol. Acad. Sci. Hung. 35, 109-l 16. Kremser P. C. and Gewertz B. L. (1985) Effect of pentobarbitol and hemorrhage on renal autoregulation. Am. J. Physiot. 249, F356F360. Lawson H. and Rehm W. S. (1943) The relative value of various fluids in replacement of blood lost by hemorrhage, with special reference to the value of gelatin solutions. Am. J. Physiot. 140, 431-438. Lo&r M. (1961) The effects of splenectomy on the red blood cells of the dog with particular emphasis on the reticulocyte response. Proc. Sot. exp. Biol. Med. 107, 972-985. Lord B. I. and Murphy Jr M. J. (1973) Hematopoietic stem cell regulation. II. Chronic effects of hypoxic-hypoxia on C.F.U. kinetics. Blood 42, 89-98. Lucas A. M. and Jamroz C. (1961) Arias of Auian Hemarotogy. Agricultural Monograph U.S.D.A., Washington, D.C. Marks L. J., King D. W., Kingsbury P. F., Boyett J. E. and Dell E. S. (1965) Physiological role of the adrenal cortex in the maintenance of plasma volume following hemorrhage or surgical operation. Surgery 58, SlO-5i7. Natt M. P. and Herrick C. A. 119551 The effect of cecal . coccidiosis on the blood cells of domestic fowl. Poultry sci. 34, 1100-l 105. Naylor B. A., Welch M. H., Shafer A. W. and Guenter C. A. (1972) Blood affinity for oxygen in hemorrha~c and endotoxic shock. .I. appt. Physiol. 32, 829-833. Neuberger A. and N&n J. -S. F. (1951) Hemoglobin formation in rabbits. J. Phvsiot. 112. 292-310. Nishimura H., Nakamura Y:, Sumer’R. P. and Khosla M. C. (1982) Vasopressor and depressor actions of Angiotensin in the anesthetized fowl. Am. J. Physiot. 242, H314H324. Nouwen E. J., Decuypere E., Kuhn E. R., Michels H., Hall T. R. and Chadwick A. (1984) Effect of dehydration, hemorrhage and oviposition on serum con~ntration of vasotocin, mesotocin and prolactin in the chicken. J. I

Endocrinol. 102, 345-35 I.

Oski E. A. and Gottlieb A. J. (1971) The interrelationships between red blood cell metabolites, hemoglobin and oxygen-equilibrium curves. Prog. Hemalol. 1, 3347. Pirkle J. C. Jr and Gann D. S. (1976) Restitution of blood volume after hemorrhage: role of the adrenal cortex. Am. J. Physiot. 230, 1683-1687.

Ploucha J, M. and Fink G. D. (1986) Hern~yn~i~ of hemorrhage in the conscious rat and chicken. Am. J. Physiot. 251, R846850.

Ploucha J. M., Scott J. B. and Ringer R. K. (1981) Vascular and hematologic effects of hemorrhage in the chicken. Am. J. Physiot. 240, H9-H17. Radke W. J.. Albasi C. M. and Harvev S. (198% Hemorrhage and’ adrenocortical activity h thd fowl (Gallus domesticus). Gen. camp. Endocrinol. 60, 204-209.

Rich I. N., Heit W. and Kubanek B. (1982) Extrarenal erythropoietin production by macrophages. Blood 60, 1~7-1018.

Rocchini A. P., Gallagher K. P., Botham M. J., L.emmer J. H., Szpunar C. A. and Behrendt D. (1985) Preven-

541

tion of fatal hemorrhagic shock in dogs by pretreatment with chronic high-salt diet. Am. J. Physiot. 249, H557-H584. Rosse W. F. and Waldmann T. A. (1966) Factors controlling erythropoiesis in birds. Blood 27, 654-661. Samarut A. B. (1978) Isolation of an erythropoietic stimulating factor from the serum of anemic chicks. Exp. Cell Res. 100. 245-248. Schalam 0: W. (1965) Veterinary Hematology, pp. 335-351. Lea and Febiger, Philadelphia. Schindler S. L., &ldersleeve $ P., Thaxton J. P. and McRee D. I. (1987a) Blood volume recovery in hemorrhaged Japanese quail. Camp. Biochem. Physiot. submitted. Schindler S. L., Gildersleeve R. P., Thaxton J. P. and McRee D. I. (1987b) Hematological response of hemorrhaged Japanese quail after blood volume replacement with saline. Camp. Biochem. Physiot. in press. Shadduck R., Howard D. and Stohlman Jr. F. (1968) A difference in erythropoietin production between anemic and hypoxic mice. Prac. Sot. exp. Biol. Med. 128, 132-136. Shadduck R., Kubanek B., Porcellini A., Ferrari L., Tyler W. S., Howard D. and Stohlman Jr F. (1969) Regulation of erythropoietin XXIV. Studies on the post-hypoxic “rebound” phase. Blood 34, 477487. Siegel H. S. (1971) Adrenal, stress and the environment. World’s Poultry Sci. J. 27, 327-349.

Smith N. and l%gelbert V. E. (1969) Erythropoiesis in chicken oerinheral blood. Can J. 2001. 47. 1269-1273. Smith J. E.-and Agar N. S. (19’75)The effect df phle~tomy on canine erythrocyte metabolism. Res. Vet. Sch. 18, 23 l-236. Smith D. W. E. and Weinberg W. C. (1981) Transfer RNA in reticulocyte maturation. Bioch. Biophys. Acra 655, 195-198.

Stino F. K. R. and Washburn K. W. (1970) Responses of chickens with different hemoglobin genotypes to phenylhydrazine-induced anemia. Poultry Sci. 49, 101-l 14.

Stohlman Jr F. (1959) Observations on the physiolo~ of erythropoietin and its role in the regulation of red cell pioduciion. Ann. N. Y. Acad. Sci. 77; 71&724. Stohhnan Jr F. (1961a) The use of Fer’ and Crs’ for estimating red cell production and destruction: An interpretive review. Blood 18, 236-250. Stohlman Jr. F. (1961b) Humoral regulation of erythropoiesis VII. Shortened survival of erythrocytes produced by erythropoietin or severe anemia. Proc. Sot. exp. Biol. Med. 107, 884-887. Sturkie P. D. (1943) The reputed reservoir function of the spleen of the domestic fowl. Am. J. Physiot. 138,599--6@2. Sturkie P. D. (1976). Asian Physiology, pp. 53-75. Springer, New York. Swan H. (1965) Experimental acute hemorrhage: The relation of blood pressure change to plasma dilution. Arch. Surg. 91, 390406. Swingle W. W., Kleinberg W. and Hays H. W. (1944) A study of gelatin and saline as plasma substitutes. Am. J. Physiot. 14, 329-337.

Szlyk P. C., King C., Jennings D. B., Cain S. M. and Chapler C. K. (1984) The role of aortic chemoreceotors during acute anem&. Can. J. Physiot. Pharmacoi 62, X9-523.

Torrence J., Jacobs P., Restrepo A., Eschbach J., Lenfant C. and Finch C. A. (1970) Intraerythrocytic adaptation to anemia. N. Engl. J. Med. 283, 16>168. Vandecasserie C., Schnek A. G. and Leonis J. (1971) Oxygen-affinity studies of avian hemoglobins. Eur. J. Biochem. 24, 284-287.

Wilson J. X. and Butler D. G. (1983a) Adrenalectomy inhibits no~d~ner~c, adrenergic and vasopressor responses to angiotensin II in the Pekin duck (Anas ptaryrhynchos).

Endocrinology 112, 645-652.

542

S. L. SCHINDLER and R. P. GILDERSLEEVZ

Wilson J. X. and Butler D. G. (1983bj ~~ydroxydo~amine treatment diminishes noradrenergic and pressor responses to angiotensin II in adrenaiectomized ducks. Endocri~otogy 112, 653458. Wyse D. G. and Nickerson M. (1971) Studies on hemorrhagic hypotension in domestic fowl. Can. J. Physioi. Pharmacot. 49, 919-926.

Younes R. N., Aun F., Tomida R. M. and BiroIina D. (1985) The role of lung innervation in the hemodynamic response to hypertonic sodium chloride solutions in hemorrhagic shock. Surgery 95, 900406. Zambraski E. J. and SchuIer R. (1980) Failure of prostaglandin inhibition to attenuate the tolerance to hemorrhage in domestic chicken. Poultry Sci. 59, 2567-2569.