. . ... _ . . 35...... ...... ...4.....__:......_.. .. . . .. \_ ....... :2 . I.............4... .. .. ... ... ... ..._. .. ' ... . ... ...-.. .14... .4... .. 4.4.44 ...44 4........4...4. .. . .. .. ..wfi..............4 4.... .4 .... ..v. .44........._.4.......4..4._.444.4........ 4 .. 4.... ... z . ....-.) .4......4..._.... .......L....../...44.. 34.42.432.44 ......4......4...4......._ .....4.......... .......4...._..“__....444. ..44 . 444.44.44.44 .4 ...... ......4 ...:4 . .45..4.~ ..........4... . .. ......L. 1 .. _ .. ......I...... .....4...4... . ...; 1.2.4.2.. ................... .... 4... .. -.. .4. . 4.2.1.4... ._ . .... 2.3.14.1: 33...... .. ....4... 4.... ...... 474.4 4a... .... .. . . ._ . ... . ........._...._.._. . . .. ............._.._. 4...... ........4..._ 4..4 ...... . ... . .... ... 4.5.... .0 umhnr: o 4—3 v .x—l‘.~H_ 24‘93a4a’4‘4ar 44.4.5444... .41.... . _ 4.4... ..4..,.4..4..._ ...r....4..... ..f 4. ...4... 4. . . .4... 44 4.. ...—.44 43.44... ..4._....4........_ ... . _. :44... 4......4..4.... .... ......4..._... .4 .... .4......... ...4 .4 3.... ... . ..u..,4~_4._. 4 ..J .41....44...n4 ... 3.4.4.4.... .... .c....1.......... . . .4. 3.4444... 4.42.” ”4...: 4......._..H44. ..4 44.... ... ...... 4 . .....H...4.... . ”£432.... . 444.44... . _ .....2 4.4 I... .... ...4. .44......; 4......4.4. 4... . ”4.144... 44.....3...:... 1.14:... .44 a... ......... .... ..4..... £24... . . 1.4.4.4....441144 4.. . ....44........ 44.44.... 4 .4.44. 4 ........4.......... . . 4... ...... 4.1.... ...... ........_..4....._.:. .....2..-..... .44 a. ......r . 4 _ A a .....m..........,... ...... .41.... ...... 44.44.24.411. 4...... .41.. 4...». ........... .....J. ......... .....- ............ ......4..... ...: .. 2.3.... 1...... . . . .. .. .... . 4 . . . r. I '4; . . 4.1- 4 :44. .v 4. . . . ..44,_4................... (.4... . . .. , ..4.........-..:.. .. . . .. . . . .......,........._. . . . . , . , . .. . . . . ”.Inuio . . . . . , .14 ......41...44.....4.., . . . . . . . . . ... . . . . ......44: . .3.” “4. . . , , . . . . ...4...._. . . ..........u....... 4 . . v 4 4V :— A a r. l . . ‘ .44nl.ain. 4 ... 4..."... .. . . .1... ... . ... .. 37.5.71... , . ......f...... . ... . . . . . . .r . . . . . . . . . 2V. . . . ...........4.|. . . . .... _ . . . ......r...............un4...44..4.fl.v..4b....u .. .. T ... . .... . . ...: . . ... . I v ....zn: . . . .. . 1:. . .i . ... . ... .... .... ...: 3...... . 4 . . .. r. . ...... .2 ...... .. $453.. .... 4,1444 .... , .. ... .... 4..L..._....;4.v.4 ... . . . ... .. . .... .. . . .. . . 4.7.5... ...—.4. . .24.. F.34WM-SW1VE-4wm .........:.. ..44.4.~4‘.44‘..—e x...— 4.... ..I_4 ... ...u...i 'l‘huutocerufgdmflm thesis entitled _ r; ". EFFECT OF ACUTE METABOLIC ACIDOSIS 0N RENAL GLUCOGENESIS presented. by 5 Raymond Lucas Burich \ has been accepted towards fulfillment of the requirements for Phone degree in PhYSiOIOgY Major professor Date 9/17/70 ' 0-169 ABSTRACT EFFECT OF ACUTE METABOLIC ACIDOSIS ON RENAL GLUCOGENESIS BY Raymond Lucas Burich Net renal glucose production was studied in the in EEEE pump—perfused dog kidney at normal blood pH, during acute whole body metabolic acidosis, and during acute local renal metabolic acidosis. Degradation of C14- glutamine by the dog kidney was also studied to determine if glutamine can be a substrate for renal gluconeogenesis. To study the effect of acute metabolic acidosis upon net renal glucose production, dogs with in situ pump- perfused kidneys were made acutely acidotic by intravenous administration of hydrochloric acid. Renal venous-arterial blood glucose concentrations were measured (using the glu- cose oxidase-peroxidase technique) during an initial con- trol pH and after one hour of acidosis. Acute whole body acidosis caused a statistically significant change from a net renal glucose uptake during the initial control period to a net renal glucose production after one hour of acid- osis. However, in a different group of dogs, when only the renal arterial blood was made acidotic and the systemic Raymond Lucas Burich blood kept at a normal pH, the net glucose production mea- sured during the initial control period was unchanged by one hour of local acidosis. Neither whole body nor local metabolic acidosis affected renal glycogen concentrations. The data suggest that factors other than renal pH may be involved in the regulation of renal gluconeogenesis dur— ing acute changes in the body's acid-base state. 14 Randomly labeled C ~glutamine was added to whole blood and the blood incubated at 37° C for 5 minutes to 14 determine the distribution and metabolism of C -glutamine in whole blood. After 5 minutes of incubation, no C14— glutamine had entered the blood cells and only a maximum of 7.0% of the Cl4 14— —glutamine had been degraded. No C glucose was formed during this incubation. The renal degradation of glutamine was studied by injecting Cl4—glutamine into the renal artery of an in situ pump-perfused dog kidney. Immediately after the in- jection had been given, 5 consecutive one-minute total renal venous blood samples were collected and the kidney taken from the body and frozen. Cl4—amino and Cl4-organic acids were identified in each blood sample, the kidney tissue, and the injection solution by gradient elution ion exchange chromatography. The blood samples and the kidney tissue were assayed for Cl4-glucose by its conversion to Cl4—gluconic acid with glucose oxidase. The Cl4-gluconic acid thus formed from Cl4-glucose was separated from the Raymond Lucas Burich samples by gradient elution ion exchange chromatography and its total C14 activity determined. The data from 2 dogs at normal blood pH and one dog after one hour of whole body metabolic acidosis were pooled. In the renal venous blood no Cl4-amino or C14- organic acid was added by the kidney in amounts detectable by the techniques employed (techniques were sensitive to 0.10% of the Cl4 activity injected). However, a signifi- cant amount of Cl4-glucose was found in the blood. The total amount of Cl4-glucose added to renal venous blood was greater than the total amount of any single Cl4-amino or Cl4—organic acid added to renal venous blood. Kidney l4 tissue contained C activity in glutamine, glutamic acid, aspartic acid, succinate, malate, and citrate and/or pyru- vate as well as Cl4-glucose. It cannot be definitely con- cluded that absolutely no Cl4—labeled amino or organic acid produced by the kidney was added to the renal venous blood because of the limited sensitivity of the techniques. However, the data do show that the glutamine extracted by the kidney can contribute carbon atoms to renal gluconeo— genesis and that the glucose formed can be released into the renal venous blood. EFFECT OF ACUTE METABOLIC ACIDOSIS ON RENAL GLUCOGENESIS BY Raymond Lucas Burich A THESIS Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Physiology 1970 Gc7/7/ ACKNOWLEDGMENTS I would like to express my gratitude to Dr. Burnell Selleck, my major advisor, for his guidance and technical assistance during my graduate program. I would also like to thank Dr. R. Anderson, Dr. G. Riegle, Dr. E. P. Reineke, and Dr. J. Scott for their counsel and suggestions. Special thanks are extended to my wife, Nancy, who has encouraged me throughout my doctoral program. ii TABLE OF CONTENTS ACKNOWLEDGMENTS. . . . . . . . . . . . . LIST OF TABLES . . . . . . . . . . . . . INTRODUCTION. . . . . . . . . . . . . . LITERATURE REVIEW . . . . . . . . . . . . I. The Importance of Glutamine to the Renal II. III. IV. Ammonia Buffer System . . . . A. The Renal Ammonia Buffer System. . . B. Glutamine as Substrate for Renal Ammonia Formation . . . . . . . C. Effects of Acidosis. . . . . . . Renal Gluconeogenesis . . . . . . . A. In Vivo Studies . . . . . . . . B. In V1tro Studies. . . . . . . C. Isolated Perfused Kidney Studies . . D. Substrates for Gluconeogenesis . . . E. Quantitative Importance of Renal Gluconeogenesis . . Relationship Between Gluconeogenesis and Ammoniagenesis. . . A. Metabolic Pathways of Gluconeogenesis and Ammoniagenesis . . B. In Vitro Studies of Gluconeogenesis and Ammoniagenesis . . . . . C. In Vivo Studies of Gluconeogenesis and Ammoniagenesis . . . Theories of Regulation of Renal Gluconeo— genesis and Ammoniagenesis. . . . . . A. PEP Carboxykinase . . . . . . . B. Glutaminase I. . . . . . . . . C. Glucocorticoids . . . . . . . . D. Fasting and Diabetes . . . . . . E. Insulin. . . . F. Adenine and Pyridine Nucleotides . . G. Effects of Acidosis on Glutamine Oxidation . . . . . . . . . . Page ii vi METHODS I. II. III. Study of Glucose Production with the In Situ Pump—Perfused Kidney . . . . . . A. Surgical Technique . . . . . . . B Blood Samples. . . . . . . C. Chemical and Physical Analyses . . Study of C14— —Glutamine Breakdown in Whole Blood. . . . . . . . . . . . A. Procedures. . . . . . . . . . B. Calculations . Study of C14- -Glutamine Metabolism in the In Situ Pump— —Perfused Dog Kidney. . . . A. Procedures. . . . . . . 1. Surgical technique. . . . . . 2 Experimental procedure . . 3 Preparation of blood and kidney tissue filtrates . . . . . . 4. Amino acid assay . . . . . . 5. Organic acid assay. . . . . . 6. —glucose assay . . . . . . B. Reagents . . . . . . l. Cl4-glutamine injection solution . 2. Hydrochloric acid solution . . . 3. Amino acid analyzer solutions . . 4. Organic acid analyzer solutions . 5. Glucose assay . . . . . . . C. Calculations 1. Total C14 activity in renal venous blood and kidney protein- free filtrates. . . 2. C14 activity in the renal venous blood and kidney perchloric acid precipitates. . . . . . 3. Renal glutamine extraction . . 4. Cl4 activity contained in each specific amino acid . . . . 5. Total C14 activity contained in amino acids . . . 6. Cl4 activity contained in each specific organic acid. . . . 7. Total C14 activity contained in organic acids . . . . 8. Total C14 activity contained in the glucose fraction . . . . . 9. Statistical analysis . . . . . iv Page 38 63 64 65 65 66 66 66 67 67 lb \— RESULTS. . . . . . . . . . . . . I. Effect of Whole Body Acidosis on Renal Glucose Production. . II. Effect of Local Renal Acidosis on Renal Glucose Production. . III. Cl4— Glutamine Distribution and Metabolism in Whole Blood . IV. Study of C14— Glutamine Metabolsim in the In Situ Pump— —Perfused Dog Kidney . DISCUSSION. . . . . . . . . . . . I. Effect of Whole Body and Local Renal Acid— osis on Renal Glucose Production . II. Cl4— Glutamine Distribution and Metabolism in Whole Blood III. Study of C14— Glutamine Metabolism in the In §£EE Pump— —Perfused Dog Kidney . SUMMARY AND CONCLUSIONS . . . . . . . BIBLIOGRAPHY . . . . . . . . . . . APPENDIX . . . . . . . . . . Frequently Used Abbreviations . . . Page 69 69 72 72 74 100 100 104 105 113 118 128 129 10. ll. 12. LIST OF TABLES Page Contents of Varigrade Chambers. . . . . . 53 Cumulative Data from the Experiments on the Effect of Whole Body and Local Renal Acidosis on Renal Glucose Production. . . . . . 82 Glycogen Content of Kidney in Control, Whole Body Acidosis, and Local Acidosis. . . . 83 Effect of Whole Body and Local Acidosis on Mean Systemic and Renal Perfusion Blood Pres— sure . . . . . . . . . . . . . . 84 Cl4—Glutamine Distribution and Metabolism in Whole Blood . . . . . . . . . . . . 85 Total C14 Activity in Protein-Free Filtrates and Precipitates from Experiment I . . . . 86 Total C14 Activity in Protein-Free Filtrates and Precipitates from Experiment II . . . . 87 Total C14 Activity in Protein—Free Filtrates and Precipitates from Experiment III. . . . 88 Cl4 Activity of the Amino Acid, Organic Acid, and Glucose Fractions in the Blood and Kidney Samples from Experiment I . . . . . . . 89 Cl4 Activity of the Amino Acid, Organic Acid, and Glucose Fractions in the Blood and Kidney Samples from Experiment II . . . . . . . 90 Cl4 Activity of the Amino Acid, Organic Acid, and Glucose Fractions in the Blood and Kidney Samples from Experiment III. . . . . . . 91 Cl4 Activity Contained in Each Amino Acid in the Blood and Kidney Samples and Injection Solution from Experiment I . . . . . . . 92 vi Table Page 13. Cl4 Activity Contained in Each Amino Acid in the Blood and Kidney Samples and Injection Solution from Experiment II. . . . . . . 93 14. Cl4 Activity Contained in Each Amino Acid in the Blood and Kidney Samples and Injection Solution from Experiment III . . . . . . 94 15. Cl4 Activity Contained in Each Organic Acid Area in the Blood and Kidney Samples and Injection Solution from Experiment I. . . . 95 16. Cl4 Activity Contained in Each Organic Acid Area in the Blood and Kidney Samples and Injection Solution from Experiment II . . . 96 17. Cl4 Activity Contained in Each Organic Acid Area in the Blood and Kidney Samples and Injection Solution from Experiment III . . . 97 18. Total C14 Activity in Renal Venous Blood Samples 1 Through 5 in Each Organic Acid Area Expressed as a Per Cent of the Cl4 Activity in the Injection Solution for That Area. . . 98 19. General Data for Experiments I, II, and III on Metabolism of C14-Glutamine. . . . . . 99 3+. $T‘."°“_.. - o ".- to ~‘ up ' . . Oust Hurmfi’rr-r. I; 77;: at: :41}:- if: ‘z INTRODUCTION Although the kidney contains metabolic pathways which are present in many other organs (glycolysis and the tri- carboxylic acid cycle), the kidney like the liver often produces glucose rather than utilizing it. Net glucose production from the kidney has been measured (in 2122) in the dog and has been shown to be a result of gluconeo- genesis (Steiner SE n£., 1968). Not only is gluconeo— genesis a metabolic pathway not usually active in other organs (except liver), but it is also one affected by the acid-base balance of the body. Thus, in the kidney there appears to be a relationship between the acid-base state of the body and at least one phase (gluconeogenesis) of renal metabolism. This relationship is explored more deeply in this thesis. The kidney has an important role in the correction of metabolic acid—base disturbances in the body. Although acute changes in the body's acid-base state may be rapidly compensated for by the removal of carbon dioxide via the lungs, chronic excesses of nonvolatile acids (sulfuric, phosphoric, and hydrochloric) must be excreted via the kidney. Excesses of the above nonvolatile acids are ex— creted by the kidney in combination with ammonia. The augmented renal excretion of ammonia during metabolic acidosis has been shown in man (Sartorius SE 21., 1949), dogs (Steiner en_gi., 1968), and rats (Rector SE n;., 1955). Since during normal acid-base balance and during metabolic acidosis the major urinary precursors of am- monia are the amide and amino nitrogens of glutamine (Pitts, 1968), this substrate has a very unique role in the kidney's function as well as its metabolism. Concurrent with early studies on renal compensatory responses to metabolic acidosis were investigations demon- strating that the kidney could contribute glucose to the systemic circulation (Reinecke, 1943; Reinecke and Hauser, 1948). Krebs later studied renal cortex slices from rats (Krebs en ni., 1963b; Krebs and Lund, 1966) and found that these slices were capable of forming glucose through gluconeogenesis. Krebs' studies have been confirmed in dog renal cortex slices (Goorno SE 21., 1967). These later studies not only showed that acidosis caused in- creased gluconeogenesis in both rat and dog renal cortex but also indicated the possibility that glutamine could be used as a substrate for the gluconeogenesis observed. With the information that acidosis stimulated both gluconeogenesis and ammonia production (ammoniagenesis), it has been subsequently postulated that the 2 systems in the kidney may be related and that glutamine is a sub— strate for both renal ammoniagenesis and gluconeogenesis. This thesis has been designed to answer some basic ques— tions relating to renal glutamine metabolism since from the above cited studies, this substrate seems to be uniquely involved in renal function during acid-base dis- turbances. In order to study renal glucose metabolism, a sensitive assay for blood glucose and an in 2129 prep- aration, where the blood flow was constant and accurately known, were developed. These techniques were utilized to: (1) study renal glucose production or uptake at normal body pH, during whole body acidosis, and during local renal acidotic conditions and (2) study the inter— mediary metabolism of Cl4-labe1ed glutamine both in the normal and acidotic dog kidney to determine whether any of its degradation products are discharged into the renal venous blood. The data presented in this thesis add to our knowledge of the effect of whole body and renal acid- osis upon net renal glucose uptake and production as well as to our knowledge of the metabolic fate of renal gluta- mine. LITERATURE REVIEW I. The Importance of Glutamine to the Renal Ammonia Buffer System A. The Renal Ammonia Buffer System During normal acid—base balance the hydrogen ion secreted by the kidney is neutralized in the renal tubule by the base derived from filtered sodium phosphate and sodium bicarbonate. During acid stress this source of base is rapidly utilized and the amount of strong acid that can be excreted in the free form is limited because at a urine pH of 4.5 hydrogen ions can no longer be secreted by the renal tubule. However, the animal can further increase excretion of acid by combining the acid with ammonia. Therefore, during metabolic acidosis, ammonia excretion increases in response to the increased need to eliminate acid and to conserve body stores of base. The ammonia buffer system is usually described as a passive system in which nonionic ammonia diffuses into the urine. Ammonia (NH3) in this system is available either as preformed ammonia in renal arterial blood or from the degradation of ammonia—containing compounds. Although the pK of ammonia is high (pK = 9.2) and at a cellular pH of 7.4 only 1% of the total ammonia and ammonium (NH4+) 4 n. I; . a! t s 3. , l 1‘: Wag-A J! l- h _ f,_ A . 9,00 . d :_5 ’ exists as the free base NH ammonia can freely diffuse 3, across the tubular cell membranes whereas the ammonium ion cannot (Jacobs, 1940). Ammonia diffusion from the blood to urine has been further demonstrated with studies in which bicarbonate, acid, or ammonia were rapidly injected into chronically acidotic dogs (Balagura and Pitts, 1962). An injection of ammonia and creatinine shows that ammonia concentration increases in the urine before creatinine appears; this finding demonstrates that ammonia moves into the urine downstream from the glomerulus. Thus, ammonia can move from the peritubular blood into the urine. Then when the dogs were made alkalotic and urine pH rose to 8.0, no ammonia appeared in the urine ahead of the creatinine. When NaHCO was given intraarterially to dogs at normal 3 blood pH (Sullivan and McVaugh, 1963), urine ammonia con- centration increased before urine creatinine appeared but as soon as NaHCO3 appeared in the urine and the urine pH rose, the urine ammonia concentration fell. Decreasing arterial blood pH caused an opposite effect. These ex— periments support the concept that: (l) ammonia diffuses readily and in both directions from the blood to tubular fluid and (2) the direction of diffusion is from the alkaline phase to the acidic phase and down an ammonia (not an ammonium) concentration gradient from blood to urine or vice versa. There is no apparent evidence in mammals that NH3 is actively transported across cell mem— branes. In the renal tubule (especially distally) where tubular pH is lower than cellular pH, ammonia binds with H+ and is trapped in the tubule as NH + which is excreted 4 in association with an anion. Most of the ammonia is added distally where the pH differential is greatest but can be added proximally (Glabman SE 21., 1963) especially during chronic metabolic acidosis (Clapp SE n1., 1965; Hayes SE 31., 1966). B. Glutamine as Substrate for Renal Ammonia Formation When humans, rats, or dogs are presented with a daily acid load, there is a gradual increase in ammonia excretion (Sartorius SE n1., 1949; Rector SE 21., 1955; Steiner SE 31., 1968) which is not due just to lowering urine pH (Pitts, 1964). The source of renal ammonia ex- cretion during metabolic acid stress and normal acid—base balance is largely from glutamine. Van Slyke SE 21. (1943) found that glutamine infusion into an acidotic dog could increase ammonia excretion but when the dog was made alkalotic with bicarbonate, renal extraction of glutamine promptly fell. This evidence led VanSlyke to suggest that glutamine might be the major urinary ammonia source since its uptake could account for 60% of the ammonia produced. Infusion of a variety of amino acids into acidotic dogs can cause increased ammonia excretion (Lotspeich and Pitts, 1947) but this technique does not define the source of ammonia either at endogenous blood levels of amino acids or at normal blood pH. Measurements of venous-arterial blood concentrations of amino acids give a more accurate picture of the amino acids extracted or released by the kidney under normal and acidotic circumstances. Owen and Robinson (1963) measured amino acid uptake and production in the human kidney by sampling brachial artery and renal venous blood. In non-acidotic human subjects glutamine, proline, and glycine are significantly extracted but only glutamine consistently in all 6 subjects. Serine, glutamic acid, alanine, cystine, and arginine were released by the kidney but only serine in all subjects. During acidosis the net extraction of glutamine increased while proline, glycine, and valine were still extracted to the same de- gree. Glutamic acid was also extracted instead of being released. Shalhoub gn_n1. (1963) carried out a similar study on dogs. During ammonium chloride acidosis, glu- tamine plus asparagine, citrulline, tryptophan, and pro- line were taken up by the dog kidney and alanine, serine, glutamic acid, cystine, and ornithine were added to renal venous blood. In these latter studies the net extraction of amino and amide nitrogens could account for 95% of the ammonia produced in acidosis, 80% in acute metabolic acidosis, but only 11% in acute alkalosis superimposed on chronic alkalosis. This finding indicates that changes in renal amino acid extraction can account for changes in renal ammonia production which occur during steady state acidosis. However, acute changes in the acid—base state may not be followed by immediate extraction changes, since renal intracellular stores of ammonia may produce con— siderable ammonia until extraction changes occur (Pitts SE 21., 1963). Glutamine is also the major precursor of ammonia in the acidotic rat (Lyon and Pitts 1n Pilkington SE 31., 1970). Glutamine is actively reabsorbed from the tubular lumen (Lyon and Pitts, 1969) as evidenced by the fact that when the lumen concentration of glutamine is near 0, tis- sue levels can be 0.344 moles/g in acidosis and 0.631 moles/g in alkalosis. But Pilkington en 31. (1970) found no Tm for glutamine reabsorption even when plasma levels were raised 10-fold. During respiratory acidosis and to a lesser degree during metabolic acidosis, they found in— creased plasma levels of glutamine in the dog. During chronic acidosis and to a lesser degree during reSpiratory acidosis there is higher ammonia production than can be accounted for by only tubular reabsorption of glutamine. This indicates passage of glutamine from the antiluminal border into the tubular cells. Whether this uptake is active or passive has not been experimentally determined but the plasma amino acid concentrations are usually less than those intracellularly, indicating an active process. During normal acid-base balance glutamine extraction may be more closely related to the quantity filtered and re- absorbed thus requiring little contribution from anti— luminal transport as suggested by Oelert and Negal (1966). By injecting N15 —labeled compounds into the renal arteries of acidotic dogs, it is possible to identify the source of urinary ammonia. Data has been compiled by Pitts (1968) from experiments on dogs showing that 43% of urinary ammonia comes from the amide nitrogen of glu- tamine, 18% from the amino nitrogen of glutamine, 6% from alanine, 4% from glycine, 2% from glutamic acid, and 35% from arterial ammonia. This data confirms earlier observations by VanSlyke 2E 21. (1943) that glutamine is the major precursor (73% of the ammonia formed in these experiments) of renal ammonia. If glutamine is the major precursor of ammonia in acidosis, then glutamine ought to be produced in increased quantities in the kidney or at some other site in the body during acidosis. Blood glutamine levels in the dog remain unchanged or increase during acidosis even though renal glutamine extraction increases (VanSlyke 2: 21., 1943; Shalhoub 2: 21., 1963). Since renal glutamine synthetase is absent in the dog (Lyon and Pitts, 1969), the kidney is unable to synthesize glutamine and must extract it from 10 renal arterial blood. The glutamine necessary to maintain blood levels comes from the liver which increases its glutamine production during acidosis (Addae and Lotspeich, 1968). C. Effects of Acidosis Slice studies of renal cortex in dog and rat have supported the whole body studies showing that metabolic acidosis stimulates increased ammonia production. If the renal cortical slices from normal rats are compared to those from acidotic ones (usually after 3 days of NH4C1 feeding) the latter slices show a higher production of ammonia (Goldstein, 1967; Goorno 2: 21., 1967; Kamm and Asher, 1970). Similar results are obtained with dog renal cortex slices (Goorno 2E 21., 1967). Goorno 2E 21. (1967) found increased ammonia produc- tion from renal cortex slices of rats with respiratory acidosis of 12 to 24 hours. After 2 days of respiratory acidosis, however, there was no longer an increase in ammonia production. There is little further information available on the effects of respiratory acidosis on am— monia excretion. In the rat the rate of urinary ammonia excretion does not seem to be limited by plasma glutamine concen- tration even though glutamine extraction increases during acidosis (Bignall 2; 21., 1968). Janicki and Goldstein (1969) found no correlation between glutamine 1.; ‘u'fi‘ 'J- 1 _ «am. .... g I I" I :‘Lfsmisa‘ci .3.) 11 concentration and renal phosphate-dependent glutaminase activity or hepatic glutamine synthetase activity. When glutamine synthetase is inhibited by methionine sulfoximine in the rat, the animal still adapts to acid loads by in- creased ammonia production and excretion (Weiss and Preuss, 1970). Thus, plasma glutamine concentration and activity of glutamine synthetase do not seem to be the only factors responsible for increased renal ammonia production during acidosis in the rat. As noted earlier when humans, rats, or dogs are pre- sented with a daily acid load, there is a gradual increase in ammonia excretion and glutamine is the major precursor of this ammonia. Thus, the 1n_y1zn and 1n‘vitro investi- gations have identified glutamine as the major precursor of renal ammonia during normal and altered acid-base bal— ance. Glutamine metabolism is therefore associated with renal ammoniagenesis. Glutamine's association with renal gluconeogenesis can also be demonstrated. II. Renal Gluconeogenesis It is known that the kidney is capable of glucose synthesis from noncarbohydrate precursors (gluconeo- genesis) and also that this metabolic activity may be coupled to the ammonia buffer system of the kidney. Renal gluconeogenesis with special reference to glutamine metabo- lism will be discussed in this section and the relation between gluconeogenesis and ammoniagenesis in Section III. 12 A. 1n Vivo Studies The first indication that the kidney could serve to maintain blood glucose levels came from studies of evis- cerated rabbits. Bergman and Drury (1938) determined the glucose requirement (the amount of glucose which must be given intravenously per unit body weight to maintain the blood sugar at normal levels) necessary for the evis- cerated and hepatectomized rabbit with and without normal renal function. Nephrectomy or stoppage of excretory function by ureteral ligation led to an increase in the glucose requirement of the hepatectomized or eviscerated animal. Their interpretation that lack of renal function by ureteral ligation was sufficient to raise the glucose requirement was subsequently shown to be incorrect in the rat by Reinecke 22 21. (1947). Phloridizinization or utereral ligation in rats did not prevent the kidney from acting as a glucose source in the eviscerated rat. How- ever, mercury poisoning which inhibited renal metabolism immediately led to hypoglycemia in their preparation. Reinecke (1942) found that the non-nephrectomized, eviscerated rat survived without added glucose longer than the nephrectomized, eviscerated rat. Both groups died of hypoglycemia suggesting that the kidney might have a role in blood glucose homeostasis. Reinecke (1943) further confirmed his theory by showing that fermentable reducing substance is added to renal venous blood (inferred to be 13 glucose) and that non—nephrectomized, eviscerated rats live up to 15 hours post-operative compared to nephrecto- mized, eviscerated rats which only live 3 hours or less. Again both groups died of hypoglycemia. Similar findings were found with the eviscerated dog (Reinecke and Hauser, 1948). It was at this time that altered metabolic condi— tions such as fasting were first shown to stimulate the gluconeogenic capacity of the kidney (Reinecke and Roberts, 1944; Roberts and Samuels, 1944). The techniques employed by these earlier investi— gators severely stressed the animal and measurements of blood glucose concentrations were not refined but the information stimulated interest in the kidney's possible role in the body's glucose homeostasis. The introduction of experiments on intact animals allowed investigators to study renal gluconeogenesis under more physiological conditions. Cohn 2: 21. (1951) used intact dogs, sampling carotid arterial and renal venous blood by indwelling catheters. Glucose was determined by the Somogyi method and a rotameter was placed on the renal vein to determine the renal blood flow. They found net uptake, production, or no venous-arterial (V—A) differences in control animals showing that renal glucose metabolism may vary from animal to animal and from time to time. MCCann (1958, 1961) using the glucose oxidase technique for glucose determina- tions demonstrated in unanesthetized dogs a rate of renal »-o . »- —_\. _ 1 _-— . I.~ " 1...’ 3“; 14 glucose synthesis which was 4—13% of hepatic glucose pro— duction. Automated refinements of the glucose oxidase technique have substantiated the data that the kidney can have net release of glucose 1n 2122 but also that it may take up glucose or show no V—A difference at all (Steiner 2: 21., 1968; Churchill and Malvin, 1970a). B. In Vitro Studies Although the intact kidney is the ideal preparation for studying renal metabolism as it exists physiologically, evidence from other preparations support the finding of 1n 2139 renal gluconeogenesis. 1n'z122 studies are indic- ative of total renal function whereas 1n XEEEQ studies can help identify metabolic patterns in cortex and medulla and allow us to study more easily gluconeogenesis from various precursors. 1n X1332 studies have shown that only the kidney's cortex can synthesize glucose (MCCann, 1962; Lee 2: 21., 1962). The medulla of the kidney is largely anaerobic and the energy necessary for medullary metabolic work (maintenance of a countercurrent multiplier) appears to be dependent on glucose degradation through anaerobic glycolytic pathways. Renal slice studies have shown that glucose can be formed from glycolytic and tricarboxylic acid cycle (TCA) intermediates as well as from amino acids. In mammalian systems there is no net conversion of fatty acid carbons 15 to glucose since they are incorporated into acetyl-CoA and subsequently oxidized in the TCA cycle producing 2 carbon dioxides and regenerating oxaloacetate (Lowenstein, 1969). However, a C14 from labeled acetyl—CoA may become incorporated into glucose because the acetyl—CoA carbons are not oxidized to CO in one turn of the TCA cycle and 2 become randomized at fumarate (Krebs 22 21., 1966). The randomly labeled oxaloacetate formed can then enter the gluconeogenic pathway. At the same time that the previously described evis- cerated rat studies were started, Benoy and Elliott (1937) incubated rat kidney cortex slices in pyruvate and lactate finding increased glucose after 1 1/2 hours indicating cortical glucose production from these precursors. Glu— cose has been shown to be produced in rat cortical slices during incubation with: glycerol, pyruvate, lactate, oxaloacetate, a—KG, glutamine, glutamate, or no exogenous substrate (Benoy and Elliott, 1937; Teng, 1954a; Goodman En 21., 1966; Krebs 2n_21., 1966; Kamm 22 21., 1967; Goorno, 1967; Preuss, 1969). Krebs and Lund (1963) and Krebs 22 21. (1963a) give a long list of other substrates and combinations including: lactate, pyruvate, malate, oxoglutarate, glycerol, succinate, citrate, acetoacetate, glutamine, proline, ornithine, fructose, galactose, man- nose, sorbose, arabinose, xylose, myo—inositol, iditol, sorbitol, xybitol, ribitol, methylglyoxal, Q . V. :4 ~ d‘eio -...x) k "1- .~ - _‘r,.5 . 4.5.14r-7 i. 16 dihydroxyacetone, glycerate, and glyceraldehyde. From this list of precursors it can be seen that the rat renal cortex has diverse metabolic machinery. Renal cortex slices from dogs have also been studied but not to the same extent as those from rats. Canine renal cortex slices will also produce glucose with no sub— strate or with glutamine or succinate incubation (Goorno 22 21., 1967; Churchill and Malvin, 1970a). It is reason— able to assume that dog cortex slices are similar to rat cortex slices for utilization of substrate although this has not been completely tested. C. Isolated Perfused Kidney Studies Limited research has been conducted with 1n Vitro perfused rat kidneys. These preparations are physiologi- cal for short periods of time (1-2 hours) with glomerular filtration and tubular functions usually at normal levels. As in the slice studies, gluconeogenesis could be sup— ported by various precursors: glycerol, pyruvate, suc— cinate, malate, fumarate, glutamate, aspartate, glutamine, and lactate (Nishiitsutsuji—Uwo 22 21., 1967; Bowman, 1970). D. Substrates for Gluconeogenesis Although the above mentioned substrates stimulate gluconeogenesis in renal cortex slices, this is not l7 absolute evidence that they are the substrates furnishing the glucose carbon atoms. Studies using C14—labe1ed sub— 14 labeling in 14 strates in rat kidney slices have shown C glucose from C14—pyruvate, Cl4-alanine, and C —glutamate (White and Landau, 1966). Cl4—acetoacetate can also pro- duce Cl4—glucose in rat cortex slices because the labeled carbons of acetyl—CoA appear in oxaloacetate which is an intermediate in the gluconeogenic pathway as well as the tricarboxylic acid cycle (Krebs 22 21., 1966). Infusion of Cl4—labeled d-ketoglutarate, malate, or glutamine into the renal artery of the intact dog causes release of C14- glucose from the kidney (Roxe 22 21., 1970). This evi— dence from Cl4-labeled precursor studies is more direct that substrate loading experiments and shows that these compounds actually contribute carbon atoms to glucose in the kidney. E. Quantitative Importance of Renal Gluconeogenesis 1n vitro, 1n 2122, isolated kidney perfusion, and isotope techniques offer strong evidence that the dog and rat kidneys are capable of synthesizing glucose from a variety of precursors. The quantitative importance of renal glucose production, however, is not so easily assessed. During long fasts (5-6 weeks) in humans, the kidneys may contribute up to 45% of the glucose utilized by the body (Owen 22 21., 1969), but its contribution 18 under normal physiological conditions is not certain. Even though the rate of gluconeogenesis (glucose formed/ min/g tissue) may be higher in kidney than in liver under some conditions (Krebs, 1963), the problem of quantita— tion of renal gluconeogenesis 1n 2122 is twofold: (l) venous-arterial glucose differences are small and are often difficult to measure due to high blood flow and (2) glucose uptake or production across the kidney will vary with time and among animals. Nonetheless, quantitation has been attempted in several studies. Roberts and Samuels (1944) measuring fermentable sugar found renal vein—aorta differences averaging 3 mg/ 100 ml blood (-1 to 6 mg/100 ml) in fed eviscerated rats. No renal blood flows were measured so net production could not be calculated. Cohn 22 21. (1951) measured renal blood flows with a rotameter in intact dogs while measur— ing renal venous-arterial glucose differences. Output from a single kidney ranged from —15.9 to 109 mg/Kg body wt/hr. MCCann and Jude (1958) improved upon the previous glucose measurements by using glucose oxidase and found renal venous-arterial glucose differences of 1.9 mg/100 ml plasma (-0.23 to 4.3 mg/100 ml) for a series of 8 dogs. Dzurik 22 21. (1963) found either glucose uptake or pro- duction depending on blood glucose levels, insulin, and adrenaline in the rat suggesting several factors may in— fluence uptake or production. 19 As experimentation continued, it became apparent that V—A glucose differences were quite small and prob- ably 2 mg/100 ml blood or less. Steiner 22 21. (1968), using an automated glucose oxidase technique, measured V—A glucose concentrations in the 1n 2122 dog kidney and reduced the standard deviation for a single glucose de- termination of 0.21 mg/100 ml blood. Experimental values obtained for glucose production by these investigators were small and in some cases negative averaging 0.09 mg/ 100 ml blood (—0.64 to 0.76 mg/ml) in their control dogs. Churchill and Malvin (1970a) utilizing a different sampl- ing technique found a small glucose uptake (0.765 moles/ kg—min) in the 1n 2122 dog kidney. These investigators suggested that rapid changes in blood glucose concentra— tions could be the cause of large errors in V-A glucose studies. Roxe 22 21. (1970) noted production from the 1n 2122 dog kidney at 1.9 mg/100 ml plasma (and also uptake in some dogs) and present firm evidence that l4-glucose gluconeogenesis does in fact occur by finding C in renal venous blood after addition of Cl4 precursors even in the dogs having net glucose uptake. Whether or not renal gluconeogenesis is important to normal glucose homeostasis has not been conclusively shown. The net rates of uptake or production must be determined and the physiological factors affecting these 20 rates defined before we can discuss the kidney's role, if any, in blood glucose homeostasis. The physiological importance of renal gluconeogenesis may not be its relationship to blood glucose homeostasis, but rather its relationship to acid—base changes and alter- ations in the ammonia buffer system in the kidney. The blood glucose homeostasis may even play a subtle role in control of acid-base problems in the body. III. Relationship Between Gluconeogenesis and Ammoniagenesis A. Metabolic Pathways of Gluconeogenesis and Ammoniagenesis When the renal cortex forms ammonia from glutamine and glutamic acid and these substrates promote gluconeo— genesis, the relationship between ammoniagenesis and gluconeogenesis becomes apparent. Thus, the metabolism of glutamine and glutamic acid is important in establish— ing the relationship between ammoniagenesis and gluconeo- genesis in the kidney. Glutamine can be deamidated to glutamic acid by glutaminase I which is a hydrolytic enzyme activated by inorganic phosphate (Sayre and Roberts, 1958) and to a varying extent by organic acids (O'Donovan and Lotspeich, 1966). In the presence of sufficient DPN+ and when the product is removed, glutamic acid can subsequently be deaminated by glutamic acid dehydrogenase to form 21 d-ketoglutarate thus liberating both the amide and amino group of glutamine as ammonia. Glutamine is not exclusively metabolized by the above pathway. There are specific renal transaminases, stimu— lated by a—keto acids (Meister, 1956), which catalyze re— moval of the amino nitrogen of glutamine to form an d—amino acid and d-ketoglutaramic acid. The d—ketoglutaramic acid formed is rapidly deaminated to a-ketoglutarate by an amidase. The 2 enzymes catalyzing these reactions are known as glutaminase II. The d—ketoglutarate formed in these reactions may enter the tricarboxylic acid cycle to be metabolized and can contribute carbons to gluconeogenesis. In species where renal glutamine synthetase is present, the cat and dog are notable exceptions (Lyon and Pitts, 1969), d— ketoglutarate can also be involved in glutamine synthesis. B. In Vitro Studies of Gluconeogenesis and Ammoniagenesis Rat cortex slices have given substantial evidence that acidosis can stimulate gluconeogenesis from several substrates and that acidosis also increases NH3 production from the slices. Renal cortex slices from rats fed NH4C1 (for 3 days) when compared to control rats have been shown to have increased ammoniagenesis. These slices also had increased gluconeogenesis from endogenous substrate, _ ‘ " 29-. :fir be ”e W‘ Ho {3' -. ’ Lé,5 ”1! lzlfis 22 glutamine, glutamate, a-KG, or pyruvate (Goodman 22 21., 1966; Goldstein, 1967; Goorno 22 21., 1967). Kamm 22 21. (1967) have presented the only investi— gation of the 12 21222 effects of pH, bicarbonate, and CO2 on renal gluconeogenesis. Renal cortex slices were taken from normal rats and incubated with glutamine, glutamate, a—KG, or oxaloacetate as substrate. When the medium was made acidotic by reducing [HCO —] or by increasing CO 3 2 tension, gluconeogenesis increased. The effect could be reversed by increasing [HCO3_] or by decreasing CO ten— 2 sion. Slices from rats with respiratory acidosis also had increased glucose production. Kamm 22 21. suggested that pH rather than bicarbonate levels or CO tension is 2 the factor controlling gluconeogenesis. However, Goorno 22 21. (1967) did not find increased gluconeogenesis dur— ing respiratory acidosis. A study of the ratio (NH3 production : Glucose formed) might indicate a quantitative correlation between ammoniagenesis and gluconeogenesis. If 2 molecules of glutamine are metabolized through the glutamic acid path- way to glucose, one molecule of glucose will be formed and 4 ammonias released. If the transamination pathway is used via a—ketoglutaramate or if glutamic acid is the sub— strate, only 2 molecules of ammonia are formed for each glucose. Kamm and Asher (1970) using renal cortical slices from NH4Cl—fed rats and controls found ratios in 23 nonadx. (nonadrenalectomized) rats of 5.22 and in adx. (adrenalectomized) rats of 6.00 when glutamine was the substrate. This indicates 75% (nonadx.) and 67% (adx.) of the glutamine were converted to glucose. Ratios of 2.14 (nonadx.) and 2.96 (adx.) were obtained when glutamic acid was the substrate indicating 93% (nonadx.) and 68% (adx.) of the glutamic acid were converted to glucose. The ratios also suggest that glutamine is preferentially metabolized by the enzymes glutaminase I and glutamate dehydrogenase. The results are evidence for a correlation between in- creased gluconeogenesis and increased ammoniagenesis in— duced by metabolic acidosis. Churchill and Malvin (1970b) demonstrated that glu— coneogenesis and ammoniagenesis in rat cortex slices do not necessarily have to be coupled. Acidosis stimulated glucose and ammonia production when glutamine and gluta- mate were used as substrates, but when gluconeogenesis was partially inhibited by malonate or phenylpyruvate, ammonia production increased or remained constant. Their results indicate that the 2 pathways may be independent under certain experimental conditions but this does not exclude their mutual stimulation under physiological cir- cumstances. Research utilizing dog renal cortex slices has been limited. Goorno 22 21. (1967) compared gluconeogenesis and ammonia excretion between NaHCO3—fed and NH4C1-fed 24 dogs. Net glucose production (total glucose formed with substrate in the medium minus total glucose formed without substrate) was significantly greater in cortex slices from acidotic dogs when either succinate or glutamine were used as substrates. Administration of exogenous cortisone to the diet, which suppressed the effect of any variation in endogenous secretion of glucocorticoids, did not alter these results. There was, however, increased production of glucose from succinate, but not from glutamine, in the cortisone-treated alkalotic dogs. One interpretation of the glutamine data is that the acid-base effect is primary and not steroid—induced. In all cases, ammonia excretion increased in the acidotic animals. Churchill and Malvin (1970a) used renal cortex slices from acidotic, fasted, and control dogs and found that there was increased glucose production in fasted and acidotic dogs when glutamine was added to the bathing solu- tion. However, neither fasting nor acidosis caused in- creased ammonia production from the cortex slices indi— cating that gluconeogenesis did not stimulate ammonia production under these circumstances. These 2 papers do not warrant a conclusion on simul- taneous regulation of gluconeogenesis and ammoniagenesis in dog cortex slices but do indicate that acidosis can stimulate gluconeogenesis in dog cortex slices. 25 C. In Vivo Studies of Gluconeogenesis and Ammoniagenesis While renal cortex slices offer an easy preparation to study tissue metabolism, they are not influenced by normal blood constituents nor are they innervated. A study of any organ's metabolism is most desirable 12 2122 so that its metabolism may be studied as it exists under physiological circumstances. Only in the dog do we have information correlating gluconeogenesis and ammoniagenesis 1n vivo. Steiner 22 21. (1968) measured ammonia formation and gluconeogenesis (renal venous blood glucose - femoral arterial blood glucose) in control and NH4C1 acidotic anesthetized dogs, reporting that acidotic dogs had both increased glucose and ammonia excretion 12 2122 compared to control dogs. Since no change was observed in total renal carbohydrate content (glucose plus glycogen) of the dogs examined, true gluconeogenesis probably occurred. A quantitative analysis of the results gives a glucose to ammonia ratio of 1:3.9 which is within the expected values of 1:2 to 1:4 if glutamine and glutamic acid are metabo— lized as discussed for the 1n 21222 study of Kamm and Asher (1970). This ratio suggests a correlation between glutamine and/or glutamic acid metabolism and ammonia pro— duction as a result of common stimulation by acidosis. This does not, however, suggest that gluconeogenesis 26 controls ammoniagenesis or vice versa. This latter con— cept was tested 12 2122 by Churchill and Malvin (1970a) using control, fasted, and NH4C1 acidotic anesthetized dogs. Control dogs exhibited glucose utilization which was unchanged by acidosis whereas ammonia excretion was increased in acidotic dogs. Fasted dogs had less glucose uptake (possibly by increased gluconeogenesis) and less ammonia excretion than controls. Their data point out that gluconeogenesis and ammoniagenesis are not changing in the same direction under conditions of fasting and acidosis. IV. Theories of Regulation of Renal Gluconeogenesis and Ammoniag2nesis Regulation of renal gluconeogenesis and ammonia- genesis during alterations in acid—base state may occur by several types of mechanisms. Regulation by enzyme stimulation or 22 2222 enzyme synthesis had been suggested for the renal enzymes glutaminase I and phosphoenol- pyruvate carboxykinase (PEP carboxykinase). These en— zymes are important to both renal gluconeogenesis and ammoniagenesis and their activity can have significant effects on glutamine metabolism. There are also hormones (glucocorticoids and insulin), enzyme cofactors (adenine and pyridine nucleotides), and alterations in body metabolism (fasting and diabetes) which affect renal gluconeogenesis and in some cases ammoniagenesis. 27 The regulation of renal gluconeogenesis and ammonia— genesis is probably not due to any single mechanism but rather to a combination of several. The complexity of regulation of renal glutamine metabolism during metabolic acidosis has been discussed by Simpson and Sherrand (1969) and will be reviewed briefly later in this section. A. PEP Carboxykinase Several TCA cycle intermediates, glutamine, and glutamic acid can enhance gluconeogenesis in dog renal cortex during metabolic acidosis; in contrast fructose or glycerol do not elicit this response (Goorno 22 21., 1967). Also, stimulation of gluconeogenesis by glucagon, epine— phrine, norepinephrine, and cyclic AMP occurs with lactate or pyruvate as substrates but not when fructose or di~ hydroxyacetone was used in rat livers (Exton and Park, 1968). The above data suggest gluconeogenesis regulation occurs at least in part between the TCA cycle and glyceraldehyde—3—phosphate. The enzyme PEP carboxykinase which converts oxaloacetate to phosphoenolpyruvate is a unique enzyme of gluconeogenesis and a possible control site. Its increased activity during acidosis could direct oxaloacetate to glucose synthesis rather than to citrate. Alleyne (1968, 1969) has presented evidence in rat cortex slices that PEP carboxykinase may be a regulatory enzyme. During metabolic acidosis either by ammonium chloride or 28 acetazolamide, there are increased tissue levels of phosphoenolpyruvate in rat kidney. Levels of oxalo— acetate, estimated in these experiments by aspartate levels, were unchanged or slightly decreased. These find— ings are indicative of an increased PEP carboxykinase activity. PEP carboxykinase activity has been measured in the kidneys of control and acidotic rats and found to be in- creased during metabolic acidosis (Alleyne and Scullard, 1969). This increase occurred as early as 6 hours after feeding ammonium chloride whereas glutaminase I activity did not increase until 2 days after acid feeding and glucose 6—phosphatase showed no activity changes. The data indicate that PEP carboxykinase responds to acidosis and does so before glutaminase I. PEP carboxykinase could also regulate ammoniagenesis by decreasing tissue levels of d-ketoglutarate and glutamic acid which inhibit gluta- minase I (Goldstein, 1966) thus facilitating renal ammonia production. In fact Alleyne (1970) did find a significant correlation between 1n_21222 gluconeogenesis and ammonia production in rat cortex slices after 48 hours of ammonium chloride acidosis. PEP carboxykinase activity has also been found to in- crease during alloxan diabetes and hydrocortisone treat— ment both of which can stimulate renal gluconeogenesis (Usatenko, 1970). 29 B. Glutaminase I Gluconeogenesis could regulate ammonia production by altering tissue d-ketoglutarate and/or glutamic acid con- centrations. In both rat (Goldstein, 1967; Preuss, 1969) and dog (Steiner 22 21., 1968; Balagura-Baruch 22 21., 1970) renal a—ketoglutarate and glutamic acid concentra— tions decrease during metabolic acidosis. Glutaminase I is known to be inhibited by glutamate (Goldstein, 1966; Sayre and Roberts, 1958) but the effect of a-ketoglutarate on glutaminase I is not as clearly defined where activa- tion (O'Donovan and Lotspeich, 1966), no change (Krebs, 1935), or inhibition (Goldstein, 1966; Balagura—Baruch 22 21., 1970) have been reported. Removal of a— ketoglutarate and glutamic acid because of increased glu- coneogenesis during acidosis could release the inhibition of glutaminase I and allow increased ammonia production. In the dog, supporting this hypothesis, infusion of d— ketoglutarate into acidotic dogs causes increased a— ketoglutarate and glutamic acid tissue concentrations while decreasing total ammonia production (Balagura— Baruch 22 21., 1970). Regulation at the glutaminase I level may not be the same for rat and dog, however. In rat the total renal ammonia production is a balance between production from glutamine and ammonia fixation in glutamic acid and glutamine (Damian and Pitts, unpublished observations 12 30 Balagura-Baruch 22 21., 1970). Dog kidney does not con- tain glutamine synthetase (Lyon and Pitts, 1969) and it has been suggested that 12 2122 renal production of glutamate from a—ketoglutarate (without added a- ketoglutarate) does not occur at an appreciable rate in chronic metabolic acidosis through the glutamate dehy- drogenase pathway. This is indicated because N15H4Cl injected into the renal artery of acidotic dogs does not appear in glutamic acid (Stone and Pitts, 1967), whereas when Cl4-d-ketoglutarate is injected into the renal artery, the glutamic acid pool becomes rapidly labeled. The im- plication here is that glutamate is formed through trans— aminase pathways and the ammonia is transaminated from amino acids. Thus, in the dog in contrast to the rat the renal ammonia production appears to depend largely on its rate of formation from glutamine. Glutaminase I activation or repression does not seem to be the same in all species either. During acidosis, renal glutaminase I activity has been found to increase in rats (Davies and Yudkin, 1952; Goldstein, 1965), guinea pigs (Goldstein and Kensler, 1960) but not in dogs (Rector and Orloff, 1959; Pollack 22 21., 1965) or humans (Pollack and Mattenheimer, 1965). In rats the enzyme response to acidosis can be blocked by actinomycin D without inhibiting the increased ammonia production (Goldstein, 1965). The increased activity was de novo synthesis of enzyme but even 31 when 22 n222 synthesis was inhibited, ammonia production was not impaired. The nonadaption of glutaminase I in the dog might be explained by the higher activity normally found in dogs compared to rats (Richterich and Goldstein, 1958). Possibly adaptation in the dog (increase in enzyme activity) need not occur even when the acid load is in- creased. The same has been suggested for human kidney glutaminase I and glutamate dehydrogenase (Mattenheimer 22 21., 1970). Since neither renal ammonia production nor activa- tion of glutaminase I is the same in all species, the role of glutaminase I in regulation of renal gluconeogenesis and ammoniagenesis may be species—dependent. C. Glucocorticoids Research concerning the effects of glucocorticoids has been largely on liver slices with fragmentary studies on renal tissue. The studies are important because acid— base imbalance often involves changes in glucocorticoids or an imbalance of glucocorticoids can alter acid—base balance. Renal cortex slices from normal, adrenalectomized, and glucocorticoid—treated rats have been compared with respect to glucose formation. Cortex slices from adrenalectomized rats, when pyruvate was the substrate, show decreased glucose production (Russell and Wilhelmi, 32 1941). Cortisol, dexamethasone, and triamcinolone treat- ment causes increased incorporation of Cl4 14 into glucose from Cl4-pyruvate, Cl4-alanine, or C -glutamate in rat cortex slices (White and Landau, 1966). Adrenalectomized rats often have decreased food intake compared to their normal controls which superimposes fasting on steroid effects. Since fasting is known to affect gluconeogenesis, Yoshida 22 21. (1969) pair—fed normal and adrenalectomized rats to exclude fasting effects. Under these conditions adrenalectomized pair—fed rats converted less succinate, malate, or glutamic acid to glucose in cortex slices than control rats. Administration of steroids increases renal PEP carboxykinase activity and urine ammonia levels in rats (Alleyne and Scullard, 1969). Kamm and Asher (1970) found that adrenalectomy does not significantly reduce ammonia production in rat cortex slices. Although glucocorticoids have been shown to affect renal gluconeogenesis, a regu- latory role of glucocorticoids in ammoniagenesis is not clearly established. D. Fasting and Diabetes Fasting an animal enhances its ability to synthesize glucose in liver and kidney indicating that conditions during fasting stimulate gluconeogenesis (Bondy 22 21., 1949; Krebs 22 21., 1963b). Diabetes also has similar effects (Bondy 22 21., 1949; Teng, 1954b; Landau, 1960) 33 where the rat kidney can incorporate Cl4—pyruvate into Cl4—g1ucose at 6 times the normal rate (Flinn 22 21., 1961). Fasting and diabetes are states in which there is not only increased gluconeogenesis, but also increased lipid metabolism and ketoacidosis. As a result of these conditions, both gluconeogenesis and ammonia production might be expected to increase. Kamm and Cahill (1969) showed that gluconeogenesis increased in rat renal cortex slices from starved or alloxan—diabetic rats but not from similar animals fed sodium bicarbonate. Churchill and Malvin (1970a) found increased gluconeogenesis in the 1n 2122 dog kidney dur- ing fasting but lowered ammonia excretion. Since the dogs were fasted only 72 hours and blood pH's were still normal, it would appear that no ketoacidosis had de— veloped. They also found increased glucose production and no changes in ammonia production in renal cortex slices from fasted dogs compared to controls. From these data, it seems that fasting affects gluconeogenesis but not ammoniagenesis, at least when ketoacidosis is absent. E. Insulin While diabetes stimulates gluconeogenesis, its counterpart (insulin availability) seems to inhibit gluconeogenesis. Reports have shown depressed rates of glucose formation with insulin (DeMeutter and Shreeve, 34 1963; Mortimore, 1963; Nadkami and Chitnis, 1963) and suppression of gluconeogenic enzymes (Weber 22 21., 1965). Insulin's effect could be caused indirectly by its anti— 1ipolytic effect on adipose tissue causing increased fix— ation of free fatty acids into lipids (Jungas and Ball, 1963; Mahler 22 21., 1964; Friedmann 22 21., 1967). Com- plicating the effect of insulin is the fact that the kidney degrades insulin (Mirsky and Broh-Kahn, 1949). If insulin degradation by the kidney is blocked by alloxan in rats, kidney slices enhance their utilization of glu- cose and pyruvate compared to non—treated animals (Mahler and Szabo, 1968). It is doubtful that insulin exerts a direct metabolic effect on the 1n 2122 kidney but rather has only indirect effects through fatty acids. The case for insulin's role in renal metabolism is still pertinent, however, when we consider its indirect effects especially during fasting and diabetes. As in— sulin levels decrease during these 2 conditions, there is increased utilization of lipids for metabolic fuel causing ketoacidosis and increased levels of acetyl-COA. The effect of acetyl—CoA on renal gluconeogenesis can be ex— plained by 2 mechanisms. It might "spare" glucogenic substrates for gluconeogenesis rather than being used for respiration, or it may activate pyruvate carboxylase which converts pyruvate to oxaloacetate (Krebs 22 21., 1965). Either or both of these mechanisms may be involved when 35 fatty acids and ketone bodies (present in greater abund- ance during fasting or diabetes) stimulate renal or liver gluconeogenesis (Krebs 22 21., 1965; Soling 22 21., 1968; Williamson 22 21., 1969). Some studies contest, however, that free fatty acids are the intermediates responsible for stimulation of gluconeogenesis during fasting in liver (Schimmel and Knobil, 1969; Exton and Park, 1967; Schimmel and Knobil, 1970). F. Adenine and Pyridine Nucleotides Tissue levels of ATP, ADP, and AMP can influence gluconeogenesis in kidney cortex. Addition of ATP or ADP to the medium bathing rat cortex slices inhibits gluconeo— genesis because of rapid conversion to AMP and the inhibi- tion is proportional to the AMP level (Weidemann 22 21., 1969). AMP inhibits fructose 1, 6-diphosphatase (a glu- coneogenic enzyme) and deinhibits phosphofructokinase (a glucolytic enzyme). Other gluconeogenic enzymes are also sensitive to the ratio of ATP:ADP:AMP. As the relative concentration of ATP is decreased and AMP increased with ADP constant: (1) conversion of pyruvate to PEP decreases, (2) PEP carboxykinase activity decreases, and (3) fructose 1, 6—diphosphatase activity decreases (Patrick, 1968). Thus, the ratio ATP:ADP:AMP can regulate several important steps of gluconeogenesis. 36 During acidosis, the degradation of glutamate is enhanced and tissue levels of glutamic acid decrease. If glutamic acid is in fact a regulator of glutaminase I, the decreased tissue concentrations of glutamic acid could enhance ammonia production. Conversion of glutamic acid to d-ketoglutarate by glutamate dehydrogenase uti— lizes oxidized coenzyme. Glutamate dehydrogenase then might be regulated by the ratio of oxidized coenzyme to reduced coenzyme. Preuss (1968), in studies on rat kidney, showed that addition of either NAD or NADP to a flask con— taining rat kidney cortex mitochondria producing ammonia from glutamate could cause increased ammonia formation. Increases in the NAD/NADH ratio, which they found during acidosis, might regulate glutamate dehydrogenase and ultimately ammonia production. G. Effects of Acidosis on Glutamine Oxidation The mechanism by which acidosis stimulates renal glutamine metabolism is not fully understood and its com- plexity may be due to acidotic effects at several cellular sites. Simpson and Sherrard (1969) studied Cl4-glutamine 1 oxidation to C 40 during acidosis in renal cortex slices 2 and found that acidosis may affect several cellular mechanisms. Renal cortex slices and mitochondria were taken from dogs at normal blood pH and made acutely aci- dotic by varying medium pH and bicarbonate concentrations. 37 When medium pH and bicarbonate concentrations were de- creased, there was increased oxidation of glutamine- U—14C to C1402 in both the cortex slices and in mito- chondria. Also cortex concentrations of glutamic acid and d—ketoglutarate decreased (but only d—ketoglutarate decreased in mitochondria). These changes parallel the decreased tissue concentrations of glutamic acid and d- ketoglutarate which occur 1n 2122 during metabolic acidosis (Alleyne, 1968). The data indicate that acute acidosis may stimulate either transport of glutamine into the cells and mitochondria and/or stimulate the degradation of glutamine. When the rate of glutamine oxidation was measured in the same manner using cortex slices and mitochondria from chronically acidotic dogs, only glutamine oxidation in the slices was enhanced. The effect appears to be only cytoplasmic (possibly by increased gluconeogenesis removing the degradation products of glutamine). Whether conversion of glutamine-U-14C to C1402 is an accurate indicator of glutamine's participation in gluconeogenesis is not certain, but these data and the effects of acute acidosis on slices and mitochondria indicate that acidosis may affect glutamine metabolism in several ways. Also, the effects of acute and chronic acidosis may be different. METHODS The research performed for this thesis can be best described by segregating it into 3 sections, each with its own experimental design and method. Thus, for clarity the methods utilized in each experimental section will be dis- cussed separately under the headings: I. Study of Glucose Production with the 1n 2122 Pump-Perfused Kidney, II. Study of Cl4—G1utamine Metabolism in Whole Blood, and III. Study of Cl4-G1utamine Metabolism in the In Situ Pump-Perfused Dog Kidney. I. Study of Glucose Production with the In Situ Pump—Perfused Kidney Studying blood glucose production or uptake with the pump-perfused kidney enabled the simultaneous sampling of renal arterial and venous blood and the accurate determina- tion of renal blood flow. Knowing the venous—arterial (V—A) glucose concentration difference, the net uptake or production of glucose could be determined by multiplying this value times the renal blood flow. Since renal V-A glucose differences are very small (Steiner 22 21., 1968), they may be enhanced by lowering blood flows. This would apply if the rate of net produc- tion is not related to blood flow, an assumption which has 38 39 not yet been proven. The lower renal blood flows and the subsequent lower perfusion pressure also decrease the glomerular filtration rate which stops urine flow. With no urine flow, there is no need to correct for changes in glucose concentration of renal venous blood due to water and/or glucose excretion. The physical setup of an 1n 2122 pump-perfused kidney therefore has the following advantages: (1) it allows simultaneous sampling of renal arterial and venous blood, (2) it enables one to study renal metabolism in a prepara- tion where only the renal arterial blood is made acidotic (local acidosis), (3) renal metabolism can be studied with- out the complication of spontaneous alterations in renal blood flow, (4) the administration of PAH is not necessary to determine renal blood flow, and (5) all the normal hormones and substrates in the blood are available to the kidney. A. Surgical Technique Male and female mongrel dogs were used weighing on the average 20 Kg. They had been obtained from a local supplier and housed by the university for 10 days to determine if distemper was present. All were healthy and were fasted approximately 12 hours before surgery although the absolute duration of fasting in each individual dog was not assayable. The dogs were anesthetized by intra- venous injection of sodium pentobarbital (30 mg/Kg) with 40 additional anesthetic administered as needed during the experiment. The left jugular vein was exposed and cathe— terized for injection of pentobarbital, heparin, and acid infusion. An endotrachial tube was inserted and the dog artificially respired to maintain blood pH near 7.40 dur- ing the control periods. The left kidney was exposed by a retroperitoneal flank incision, avoiding rupture of the peritoneum when possible. The ureter was cannulated and urine flow (if any) recorded during each blood sampling period. The ovarian (females) or spermatic (males) vein was tied off in order to prevent contamination of renal venous blood. Each renal fat pad of appreciable size was also ligated to minimize any collateral blood flow to or from the kidney. The dog was heparinized (1,000 U/2 Kg) and the renal artery was cannulated with the efferent tube from a Sigma motor pump. This procedure was completed in one minute or less and perfusion was started immediately after cannula- tion. In the case of a double renal artery, each artery was cannulated separately to minimize kidney ischemia. Blood from the previously cannulated right femoral artery was used to perfuse the renal artery. The blood flow was adjusted so that renal perfusion pressure, measured 4 or 5 inches proximal to the kidney through a rubber cuff in the perfusion tubing, was less than 100 mm Hg. This was 41 necessary because acidosis tended to increase renal resistance and raise renal arterial perfusion pressure. Systemic pressure was measured at the femoral artery; mean femoral arterial pressure was within 10 mm Hg of aortic pressure. The renal vein was exposed and a bent (120° angle) 20 gauge needle inserted through the venous wall so that its tip faced the kidney. The needle was anchored in place and subsequently used to draw renal venous blood. Renal arterial blood samples for glucose and pH analyses could be drawn into a syringe through the rubber cuff in the renal arterial tubing. The dog was then allowed to equilibrate for 30—40 minutes during which time the respiration was adjusted to give a steady blood pH. B. Blood Sampling After the dog had equilibrated, paired (simultan- eously drawn) renal arterial and renal venous blood samples were taken immediately after a blood pH had been measured. Approximately 3 ml was drawn in a period of 30 seconds. Four such samples were drawn, one every 15 minutes. Immediately prior to each collection the dead space in both sampling catheters was washed out by with— drawing 5 ml of blood. After the collection period, this blood was reinfused into the jugular vein to minimize blood loss. Then the dog was made acidotic by infusion 42 of 0.5 N hydrochloric acid, made in isotonic saline, into the jugular vein. The infusion was rapid (2 to 3 ml/min) at first to lower pH and then slower (1 to 2 ml/min) to maintain blood pH at 7.10 to 7.20. After acidosis had been established, 4 more paired samples were drawn 15 minutes apart with a blood pH measured before each paired sample was drawn. During local acidosis only renal arterial blood was made acidotic while the pH of the rest of the animal was maintained normal. In order to accomplish this, a mixing chamber made from a 125 ml erlenmeyer flask was inserted between the femoral artery and the Sigma motor pump. Acid (1.5 N HCl in saline) was then infused into this chamber (0.34 ml/min) and slowly mixed with a teflon stirring bar. The mixing chamber was necessary because addition of acid directly into the perfusion tubing caused massive renal vasoconstriction. Whole body pH was maintained at control levels by increasing minute ventilation when necessary. Isotonic saline was infused at 0.34 ml/min into the mixing chamber when control renal glucose metabolism was being measured. C. Chemical and Physical Analyses Blood pressures were recorded on a Grass Model 5 recording polygraph from a Model P23AC high pressure transducer. Bloods were drawn in air-tight syringes and 43 pH's read on a Beckman pH meter utilizing an anaerobic blood sample block at 37° C. Sample pH was periodically checked on an Astrup pH meter. Whole blood glucoses were determined by the glucose oxidase-peroxidase method using Glucostat reagents ob- tained from Worthington Biochemical Corporation (Freehold, New Jersey). The whole blood drawn from the renal artery and vein was transferred to a test tube calibrated at 2 ml and 0.03 ml of 25% isooctylphenyl ether of polyethylene glycol (Triton-X 100, Lot #801123 Calbiochem) was added to hemolyze blood cells. The sample was spun down lightly to sediment any cell debris and 0.2 ml of this supernatant was added to 1.8 ml water. After mixing, 1.0 ml, 1.8% Ba(OH)2 was added and the tube remixed. One ml, 2.0% ZnSO was added and the tube mixed before centrifuging. 4 The 1.8% Ba(OH)2 and 2.0% ZnSO reagents had been pre— 4 viously balanced to neutral pH. After centrifugation, the supernatant was separated and 2.0 m1 transferred to a new set of tubes. All blood samples were immediately carried through the above procedures and were maintained at this state until all were collected and similarly pro— cessed. The glucose concentration of this supernatant was found to be stable for at least 3 hours. Standard glucose solutions were treated as whole blood and 0.2 ml of each were precipitated as above. 44 After all bloods had been drawn and treated as above, fresh Glucostat reagent was made according to the directions published by the Worthington Biochemical Corp- oration. Eight ml of Glucostat reagent was added to 2.0 m1 of each supernatant and to 2.0 m1 of water for a blank. The addition of the Glucostat reagent was spaced at 15 second intervals. The tubes and reagent were in a 25° C water bath throughout this period. After incubation for exactly 10 minutes in the 25° C water bath, the reaction was stopped by addition of 2 drops, 4 N HCl at 15 second intervals in the same order that the Glucostat reagent had been added. Each tube was mixed after the acid was added and all were left at room temperature for 15 minutes before reading the absorbancy of the solutions in a Beckman Spec- tronic 20 at 400 mu. A standard curve was constructed using 3 standards (the standards were selected so that blood values were read between the lowest and highest standard) and the blood glucose concentrations determined using the standard curve as a reference. Glucose concen— trations were expressed as mg% in whole blood. Standard error for a blood glucose concentration of 80.0 mg% was 20.20 mg% (mean : SE, N = 10). Glucose added to whole blood and carried through the procedure was 101.5% 2 0.5% recovered. Renal blood flows were measured with a graduated cylinder immediately after the experiment using blood 45 from the femoral artery. Kidney weights were also ob- tained so that glucose production or uptake could be expressed either as mg/100 ml blood or ug/min/g tissue. After obtaining the blood samples for glucose analysis in several of the whole body (4) and local acidotic studies (6), the experimental kidney was taken from the body and frozen for glycogen determinations. Kidneys from a separate group of non-acidotic dogs were used as controls. Glycogen determinations were done on frozen kidney tissue using the method of Good 22 21. (1933). The glucose obtained from the hydrolysis of glycogen was measured by the glucose oxidase technique. A one hour KOH extraction was used instead of 20 minutes as Good 22 21. (1933) describe. Slices containing both cortex and medulla were used for glycogen determinations. Exogenous glycogen added to kidney tissue containing 100 ug/g tissue was 43% i 6.0% recovered. To correct for this lack of complete recovery of exogenous glycogen, the measured glycogen concentrations were multiplied by a factor of 2.3. Renal glycogen values are expressed as pg glucose per gram wet tissue weight. A standard glycogen solution was analyzed with each tissue sample as an internal standard. Freezing did not affect glycogen concentrations in kidney tissue. 46 II. Study of Cl4—Glutamine Breakdown in Whole Blood In order to study the metabolism of a compound taken up by an organ from the blood, it is necessary to know whether or not the compound is metabolized by the blood. If the blood rapidly metabolizes the Cl4—labeled sub— strate being fed to the organ, then the organ will be sub- jected to a different group of labeled compounds than was intended and thus the renal venous blood from the organ will contain labeled products not formed by that organ. The partition of Cl4—labeled substrates between the blood cells and plasma is also an important factor to consider when studying the utilization of such compounds by the 12 2122 kidney. Whether or not glutamine enters erythrocytes must be ascertained in order that plasma glutamine concentrations can be estimated to determine the feasibility of injecting certain quantities of C14— glutamine for renal metabolic studies. If all the labeled glutamine enters the erythrocytes, then there would be no Cl4—glutamine available for renal uptake from the plasma either by glomerular filtration or by transport at the renal tubular antiluminal border (Pilkington 22 21., 1970). A. Procedures An 12 Vitro system was designed to study metabolism and partition of Cl4-g1utamine in dog whole blood. A 50 m1 erlenmeyer flask was fitted with a stopper whereby 47 the flask could be flushed with gas. The flask was then suspended in a metabolic shaker with a water bath temper- ature of 38° C. An injection solution of Cl4 14 —glutamine was prepared by adding 0.4 m1 C —g1utamine obtained from New England Nuclear to 1.0 ml of 0.02 M phosphate buffer pH 7.35 made isotonic by the addition of NaCl (0.1 ml of this was added to 4 ml water for determining the total C14 activity of the injection solution). Blood was obtained from an anesthetized, heparinized dog (with a normal hematocrit) on which only a venous cannulation had been performed. Seven m1 of this blood was added to the flask and gassed with water saturated 95% 02—5% CO2 for 15 minutes. After the 15 minutes of gassing one m1 of the injection solu- tion was added to the blood and the mixture was shaken and gassed for another 5 minutes to correspond to the length of time necessary to complete the proposed 1n 2122 experiment. After 5 minutes of incubation, the following procedures were done in the order listed: (1) one ml of the blood from the incubation flask was added to 7.2 m1 of 6% perchloric acid, (2) one m1 of blood was added to 99 ml water, (3) several hematocrit samples of the blood were taken, (4) the pH of the incubated blood was mea- sured, (5) the remainder of the incubated blood was centrifuged to isolate the plasma, and (6) one ml of this — 48 plasma was added to 99 ml water for C14 quantitation. The experiment was done twice. The whole blood in perchloric acid was treated as described in Section III, A, 3. An amino acid separa— 14 tion and a C analysis was done on the protein—free filtrate as described in Section III, A, 4. B. Calculations Total C14 activity present in whole blood was determined from a Cl4 determination performed on the one ml blood + 99 ml water solution. The total C14 activity in the plasma was determined in the same way using the one ml plasma + 99 ml water solution. If the hematocrit obtained from blood which contained the Cl4-glutamine 14 injection solution is used, then the C activity in the plasma of one ml of blood can be compared to the Cl4 14 activity in one ml of whole blood. Thus, the C activity in one ml of plasma times (l—hematocrit) will equal the Cl4 activity in one ml of whole blood if the erythro- cytes do not take up glutamine. On the other hand, if the Cl4 activity in the plasma of one m1 of blood is less than the Cl4 activity in one ml of whole blood; this indi- cates uptake of C14 by the erythrocytes. III. Study of Cl4—Glutamine Metabolism in the In Situ Pump—Perfused Dog Kidney To study the complete renal metabolic degradation products of Cl4-glutamine, a system is needed which _ 49 allows collection of the total C14 injected into the animal. The 12 2122 pump-perfused kidney allows one to completely collect the Cl4 label injected into the renal artery perfusion circuit and thus any recirculation of the original label or labeled end products is prevented. The design was such that the Cl4 -glutamine could be rapidly injected into the renal artery, all the renal venous blood and urine could be collected for the next 5 minutes, and finally the whole kidney taken from the body and frozen after the 5 minutes of renal venous blood collection. A. Procedures 1. Surgical technique.——The surgical approach to the kidney and the renal arterial cannulation were the same as described under Section I of the Methods. In addition to these procedures already discussed, the renal vein was cannulated and the renal venous blood allowed to passively drain into the left jugular vein until the labeled substrate was injected. A Y-tube was placed between the renal vein and left jugular so that the total renal venous blood could be collected after the injection of Cl4-glutamine into the renal artery. The left jugular vein was perfused with donor blood by a Sigma motor pump at the same rate as the renal blood flow during the 5 minutes of renal venous blood collection to prevent a net loss of blood from the animal. _ 50 2. Experimental procedure.——After all cannulations were completed as described above, the dog was then allowed to equilibrate for approximately 30 minutes. Dur- ing this equilibration time the Cl4 —glutamine injection solution was prepared as described under Section B, 1. In Experiments I and II immediately before injection of Cl4—glutamine, renal V—A glucose differences were deter— mined as described in Section I. In Experiment III a purity check was performed immediately on the injection solution in preference to measuring renal V-A glucose differences. Time did not allow both a purity check and a renal V-A glucose analysis to be performed. The Cl4~glutamine injection solution was loaded into a syringe and its needle was placed into the cuff in the renal arterial perfusion tube within 3 or 4 inches of the kidney. One liter plastic bottles containing 400 ml of 2° C, 6% perchloric acid were taken from a refrigerator and the experiment proceeded in the following sequence: (1) A stop watch was started to signal the beginning of the timed sequence. (2) Renal venous blood was totally diverted to the first collection bottle (containing 400 ml cold perchloric acid) from its path to the jugular vein. Immediately after beginning this renal venous blood col— lection, a Sigma motor pump was started to infuse donor blood into the jugular vein at the same rate as the renal blood flow. (3) The injection solution was then given 51 over a period of 5 seconds and it was totally injected before the completion of 10 seconds of renal venous blood collection. (4) The total renal venous blood outflow collection was then continued in separate one-minute ali— quots for 5 consecutive minutes. (5) After the above 5 one-minute collections had been completed, the C14— labeled kidney was taken from the body, cut, and placed in acetone—dry ice. (6) The urine, if any, was collected as it appeared. 3. Preparation of blood and kidney tissue fil- trates.-—After each one-minute blood sample had been col— lected, this perchloric acid-treated blood was placed in a refrigerator until prepared further. The frozen kidney was weighed and homogenized in cold 6% perchloric acid (7 ml/g kidney) in a VirTis Model 45 Homogenizer. Total volume of each perchlorate-treated blood and kidney sample was recorded. A quantitatively measured aliquot of the homogenously mixed samples of perchlorate- treated blood and kidney was poured off and centrifuged. The resulting supernatant volume was recorded and saved for neutralization with potassium carbonate; also part of the resulting precipitate of each sample was saved and stored in a freezer. The remaining uncentrifuged perchlorate—treated sample was frozen. The perchloric acid in the sample supernatants was neutralized with 52 5 M potassium carbonate in an ice bath using 3 drops of methyl orange (0.5 mg/ml) as an indicator to determine the neutralization endpoint. The volume of the neutral- ized filtrate for each sample was recorded and these neutralized samples were frozen until further analysis could be performed. The above described preparations were performed immediately after the sample collections on the day of the experiment. 4. Amino acid assay.—-Amino acids were separated for C14 assay by ion—exchange chromatography modified from a procedure described by Piez and Morris (1960). A 133 cm column of Aminex AG 50W«X12 resin (Bio-Rad Lab— oratories Control #4915) was prepared in a glass column with surrounding water jacket maintained at 28° C. A Milton Roy mini pump (Model: 196—31) was used to force the elution solution through the resin. The eluent from the column was collected with a volumetric siphon on an automatic fraction collector (Warner-Chilcott Labora— tories, Instruments Division) in 250 test tubes contain— ing 2.8 ml of eluent in each. The gradient of Na concen- tration and pH in the elution solution was provided by a 9—chambered Varigrade Gradient Mixer (Buchler Instruments). The Varigrade chambers were filled as described in Table l. The contents of the Varigrade chambers had to be modified from these described by Piez and Morris (1960) 53 so that glutamine could be clearly separated from glutamic acid. TABLE l.—-Contents of Varigrade Chambers . . . . l Citric Ac1d Na Citrate Chamber # Bufferl (m1) (ml) Water (ml) 1 75 0 0 2 75 0 0 3 75 0 0 4 73 2 0 5 64 10 0 6 4O 10 25 7 30 10 34 8 5 52 15 9 0 71 0 1 See p. 62 of this thesis for an accurate descrip— tion of these solutions. An appropriate amount of neutralized filtrate (10 to 30 m1) plus amino acid carrier standards were forced with air pressure onto the Aminex AG SOW-X12 resin. The amount of carrier amino acids added overwhelmed any endogenous amino acids present in the neutralized filtrate (Section B, 3). The space above the resin in the glass column as well as the tubing between the Varigrade and the column's pressure cap were filled with citric acid buffer. The Varigrade was then connected to a Milton Roy mini pump and the amino acid separation commenced. 54 The eluent from the Varigrade was pumped through the resin by the mini pump at constant flow, at pressures ranging from 80 to 90 PSI. After the Varigrade was empty, Na critrate was pumped until 250 eluent samples had been collected; this assured that all amino acids had been re- moved from the resin (30 hours). The resin was then re- generated by pumping citric acid buffer until the eluent from the resin was at the same pH as the citric acid buffer (4 to 6 hours). The carrier amino acids, which had been added to the column to check the consistency of the separation and to identify the specific amino acid being eluted with each C14 peak, were qualitatively located in the eluent samples with a ninhydrin assay (Lee and Takahashi, 1966). Two- tenths ml of the fraction collector sample was added to 4 m1 fresh ninhydrin reagent and then heated for 10 min- utes in boiling water. After this 10 minute incubation, the samples were cooled in an ice bath and their optical density read on a Beckman Spectronic 20 at 570 mp (430 mu for proline and HO-proline). Cl4 activity was also determined on each sample obtained in the amino acid separation. One ml of each sample was plated onto a 25.4 mm aluminum planchet and dried under standardized conditions with heat lamps. When dry, they were counted in a Baird Atomic University II Series gas flow counter. Since the dried citrate produced 55 a significant self absorption, this self absorption was quantified in each eluate sample so that the total C14 activity eluted off the column could be accurately mea— sured and thus the per cent recovery of C14 from the column could be calculated. Peaks of Cl4 activity eluted from the resin could then be correlated with ninhydrin peaks and the labeled amino acids identified. 5. Organic acid assay.——Separation of organic acids was accomplished by ion—exchange chromatography on Dowex 1 (AG 1—X8 200—400 Mesh) anion exchange resin. 'Dowex 1 resin was converted to the acetate form by washing with concentrated acetic acid followed by water rinses until all the unbound acetic acid was washed off the resin. A 10 cm X 1.0 cm diameter column of resin was poured into a glass column and allowed to settle on a sintered-glass disc. Blood or kidney tissue neutralized filtrate (15 ml) or diluted injection solution (one ml) was added to the Dowex 1 resin in addition to 2 ml of carrier organic acid solution (described in Section B, 4). The organic acids were individually displaced from the resin by a graded eluent from a double-chambered vat. The upper chamber contained the pure elution solutions (acetic acid or formic acid) which flowed by gravity into a lower con- tinuously mixed chamber initially containing 300 ml water. The volume of the lower chamber was not greatly changed 56 during the separation. The solution from the lower chamber then flowed by gravity into the column containing the Dowex l resin. The eluent from the resin column was serially divided into 160 aliquots (10 ml each) with a volumetric siphon which sequentially emptied the samples onto a fraction collector. Elution of the fraction col— lector samples 1 to 10 occurred with water in the upper chamber, samples 11 to 39 with 0.4 M acetic acid, samples 40 to 79 with 2.0 M acetic acid, samples 80 to 129 with 1.5 M formic acid, and tubes 130 to 160 with 4 M formic acid. The chromatogram obtained by the above procedure was analyzed for glutamate, aspartate, and other amino acids by the ninhydrin assay described in Section A, 4. Malate (Hummel, 1949), d-KG (Selleck 22 21., 1964) pyru- vate and lactate (Bergmeyer, 1965) were identified by chemical methods. Fumarate was identified by its absorp- tion at 213 mu. Citrate and succinate were identified by the procedure described below. Once the elution pattern had been identified by the specific assays (except for citrate), an easier and less expensive qualitative identification was established. One—half m1 from each fraction collector sample was pipetted into a test tube and dried under low vacuum and heat. To these dried samples, 2 ml of 0.00024 M phosphate buffer (containing universal pH indicator, Will Scientific, 57 Cat. #W87048) was added. If no organic acid was present in the sample after drying, the pH of the buffer remained at 7.5 as detected by both a Beckman pH meter and the color of the universal indicator. When an organic acid was present, the pH dropped to values as low as 4.0 de- pending upon the quantity and pK of the carrier organic acid added to the column. The pH peaks corresponded well with the specific assay peaks and pH analysis was used routinely for qualitative identification of the organic acid peaks. Cl4 activity was determined by plating one m1 from each fraction collector sample onto a 25.4 mm aluminum planchet, drying, and counting in the Baird Atomic gas flow counter. The first 5 tubes from the organic acid separation contained salts and other unbound material which left a film on the planchet and hence caused Cl4 self absorption in these samples. These first 5 fraction collector samples were diluted 1:11 and recounted to de— termine the average per cent self absorption. This self absorption factor was then used to correct the counts/min in the first 5 samples. All other samples (6-160) had no significant self absorption since the elution solutions acetic and formic acid are volatile and the carrier organic acids are added only in very small amounts. The mean recoveries : SE of organic acids eluted from the Dowex l resin column using procedures similar to those 58 described above were as follows: lactic acid 100% i 2% (N=3), d—ketoglutarate 91% i 2% (N=3), succinate 96% i 3% (N=3), malic acid 103% i 6% (N=3), and citrate 107% (N=2). Recoveries of these organic acids were determined with the following assays: lactate (Bergmeyer, 1965), a—ketoglutarate (Selleck 22 21., 1964), fumarate (optical density at 213 mu), succinate (Clark and Porteous, 1964), malate (Hummel, 1949), and citrate (recovery of standard Cl4-citrate added to column). 6. Cl4—glucose assay.-—The Cl4-glucose assay was based on the specific conversion of glucose to gluconic acid by the enzyme glucose oxidase. Five m1 samples of the neutralized blood and kidney filtrates were first purified by passage of the filtrates through ion-exchange resins which removed all C14 activity in amino and organic acids so that only the uncharged Cl4-labeled molecules re— mained in the sample to be assayed. This purification was performed by (1) first passing 5 ml of neutralized fil— trate through 2 pasteur pipets each containing 1.5 ml Aminex AG 50W—X12 resin (at pH 3.0) to remove amino acids, and (2) then passing the resulting eluent through 3 pasteur pipets each containing 1.5 ml AGl-X8 resin (ace— tate form) to remove organic acids. When a 5 ml sample of the Cl4—glutamine injection solution was carried through this procedure, 99.85% of the Cl4 activity was removed. 59 This purification procedure eliminated contamination of the Cl4~gluconic acid eluted from the AGl-X8 resin (des- cribed below) with C14—pyrrolidone 5—carboxylic acid. All the eluent from the above purification pro- cedure (approximately 6 ml representing 5 m1 of neu- tralized filtrate plus water present in the purification resins) was collected in a 125 ml erlenmeyer incubation flask for conversion of Cl4—glucose to Cl4—gluconic acid. Ten ml of Glucostat reagent was added to the incubation flask and the resulting solution was incubated at 37° C for 75 minutes (standard solutions of carrier and C14- glucose, when subjected to this procedure, had a maximum conversion to gluconic acid at about 60 minutes of incu- bation). The gluconic acid formed was then separated as described below from the other substances in the incuba- tion solution on AGl-X8 resin (acetate form) by gradient elution ion—exchange chromatography. The gradient elution pattern of gluconic acid from AGl—X8 resin was determined by the following procedure. Sufficient carrier gluconic acid was added to 10 cm X 1.0 cm diameter column of resin to produce a significant pH change in the eluent samples as described in Section III, A, 5. Fraction collector samples (10.3 ml/tube) 1 through 10 were eluted with water in the upper chamber and samples 11 through 25 with 2 N acetic acid. With this procedure 14 if C -glucose was added to the column, all the C14—glucose 60 came off the‘column well before sample 15. When C14— 14 glucose was converted to C -gluconic acid by the Gluco- stat incubation described above, all the C14 -gluconic acid was eluted between sample 16 and 25. Thus, for the glucose analysis of neutralized filtrates, the first 150 ml of eluent from the resin column was accurately col— lected in a graduated cylinder and then discarded. The next 100 ml of eluent was similarly collected and assayed for C14 activity in gluconic acid. Since the second graduated cylinder contained all the gluconic acid, its total C14 activity was directly related to the total C14- glucose originally present in the neutralized blood and kidney perchlorate filtrates. The total sample collected from the purification procedure (after incubation with the Glucostat) was placed onto a 10 cm X 1.0 cm diameter column of AGl-X8 resin along with 0.01 9 carrier gluconic acid and eluted with water and 2 N acetic acid as described above. Three, one ml aliquots of the 100 ml of the gluconic acid fractions were pipeted and dried on a single 25.4 mm planchet and counted in the gas flow counter. When Cl4-glucose stand- ards added to blood perchlorate filtrates were passed through these procedures, 85% i 4.3% (N=3) of the Cl4 activity was recoverable and 79.0% i 3.3% (N=3) was con- verted to gluconic acid. 61 B. Reagents 1. Cl4-glutamine injection solution.--Randomly l4 labeled C —glutamine was obtained from New England Nuclear (Lot #459—294; 0.173 mg glutamine/2.5 ml in each of 4 vials; specific activity 212 mc/mM). The shipment of C14-glutamine arrived frozen and was maintained frozen until use. Just prior to intraarterial injection, one half ml of concentrated sodium chloride was added to 2.5 m1 (one vial) of the stock Cl4 -glutamine so that the final osmolarity was 300 mOs/L. Fifty lambda of this injection solution was added to 4.0 ml of water and frozen to be used later for determining the total C14 activity and purity of the injection solution. The pH of the injection solution was approximately 4.5; however, its glutamine molarity was so low that the blood would easily buffer the injection solution to the arterial pH. Glutamine con— centration in the solution obtained from New England Nuclear was 0.48 uM/ml. When saline was added to produce an isotonic solution, this figure was reduced to 0.38 HM/ ml, 0.39 uM/ml, and 0.40 uM/ml in Experiments 1, II, and III. These levels of glutamine in the injection solution are close to those actually present in the plasma of a non—acidotic dog (0.38 uM/ml) (Addae and Lotspeich, 1968). 2. Hydrochloric acid solution.--Dogs were made acidotic by intravenously infusing a solution of 0.5 N HCl made by diluting concentrated HCl with isotonic saline. 62 3. Amino acid analyzer solutions.-—The citric acid buffer was prepared by adding 440 g of Na citrate - 2H20, 1465 g citric acid ' H O and bring— 2 ing the final volume to 18 L. One ml of caprylic acid was added and the final pH measured was 3.00 i 0.05. For each separation, 5 ml thiodiglycol and 2 m1 BRIJ 35 solution (50 g/100 ml) was added to one liter of this stock. The 0.8 M Na citrate was prepared by adding 940.8 g of Na citrate ' 2H20 to 4.0 L of water and stored at room temperature. Ninhydrin reagent was made on the day used by add- ing 0.65 g ninhydrin to 91 m1 of special citric acid buf— fer (147 9 Na citrate - 2H20, 8.0 g citric acid in l L of water) and 156 m1 glycerol (Baker Analyzed). Baker Analyzed glycerol was chosen because it had a low reagent blank. To identify the position of elution from the resin of specific amino acids, one—half ml of a standard solu— tion of that amino acid was added to the resin. Suffi— cient amino acid was present in the standard solution (.0200 mM/ml to 0.685 mM/ml) to give a quantifiable ninhydrin reaction when analyzed in the fraction col— lector samples. Not all standard amino acids were used for each amino acid separation; only those amino acids 14 most likely to become labeled from C —glutamine were assayed with ninhydrin. 63 4. Organic acid analyzer solutions.--Two-tenths g of each of the following organic and amino acids were added to 30 ml water to make the organic acid standard: fumarate, succinate, d-KG, malate, citrate, pyruvate, lactate, aspartic acid, and glutamic acid. Two ml of this carrier organic acid solution was added to each sample of neutralized filtrate just prior to loading the sample onto the resin column so that the position of elu- tion of each organic acid could be identified. Fifty mg of pyrrolidone 5-carboxylic acid was also added directly to the sample to be separated. 5. Glucose assay.—-Aminex AG 50W—X12 resin was prepared by washing with a pH 3.00 HCl solution. Cl4-glucose was used as shipped from Calbiochem (D—glucose-U-C-l4 in 0.25 ml 25% ethanol. Lot 930099. 100 no with specific activity 50 mc/mM). C. Calculations 1. Total Cl4—activity in renal venous blood and kidney protein-free filtrates.-—Since only part of the total blood-perchloric acid mixture was neutralized, the per cent of protein—free filtrate (PFF) (the supernatant portion of the mixture) in each blood—perchloric acid mixture was calculated. The per cent of PEP in the por— tion of the blood-perchloric acid mixture centrifuged equals the volume of decanted fluid divided by the initial 64 volume before centrifugation. This per cent times the total blood—perchloric acid mixture volume equals the total volume of PFF in the blood-perchloric acid mixture. The remainder of the mixture after centrifugation was precipitate. Total C14 activity in the neutralized PFF was de- termined by making a 1:21 (Experiment I and II) or 1:11 (Experiment III) dilution of the neutralized PFF, plating one ml of the diluted filtrate on a 25.4 mm planchet, and counting in the gas flow counter. The counts/minute 14 activity per m1 of measured times 21 or 11 gives the C neutralized PFF. Several of these neutralized PFF's were further diluted to 1:121 and recounted to determine the self absorption. The values were then corrected for self absorption by the factor 1.16 (Experiments I and II) and 1.24 (Experiment III). The activity per ml of neu— tralized PFF times its total volume equals the total activity in the volume of PFF which had been neutralized. If the total unneutralized volume of PFF is divided by the volume of PFF neutralized, the ratio obtained times the total activity in the neutralized PFF will equal the total C14 activity in the total PFF. 2. Cl4 activity in the renal venous blood and kidney 14 perchloric acid precipitates.-—The C activity in the precipitate from the blood and kidney perchloric acid mixtures was estimated by dissolving one g of precipitate V 65 in 15 ml 2 N NaOH, plating one m1 of the solution, and counting in the gas flow counter. The activity was mul- tiplied by 15 to correct for volume and then by the total number of grams of precipitate. Self absorption was de— termined by adding 0.1 m1 of a Cl4 solution of known activity and repeating the procedure. Each Cl4 activity value from both blood and kidney precipitates was then multiplied by the self absorption correction factor (5.5). 3. Renal glutamine extraction.—-The glutamine ex- tracted by the kidney was the total C14 activity injected 14 activity as glutamine in the 5 renal venous blood samples. The C14 activity as as glutamine minus the total C glutamine was determined by gradient elution ion-exchange chromatography of each of the injection solutions and whole blood perchlorate filtrates. 4. Cl4 activity contained in each specific amino acid.—-The total C14 activity contained in each specific amino acid in the total PFF is determined by the product of the PFF's total C14 activity and the per cent of the PFF's Cl4 activity in that amino acid. The per cent of the PFF's Cl4 activity in that amino acid was calculated as the total C14 activity in the amino acid peak divided by the total C14 activity eluted from the resin column. The per cent recovery of C14 from the Aminex AG 50W—X12 resin during amino acid separation was 90% i 3% (N=18). 66 5. Total C14 activity contained in amino acids.—- The amino acid chromatogram began at approximately tube 14 50 of the amino acid separation. The C activity mea— sured in tubes 51 through 250 was divided by the total C14 activity in the chromatogram to give the per cent Cl4 activity as amino acids. The per cent times the total 14 activity in the PFF gives the total C activity contained in the amino acids of that filtrate. 6. Cl4 activity contained in each specific organic acid.--The total C14 activity of each specific organic acid in the total PFF is determined by the product of the PFF's total C14 activity and the per cent of the PFF's Cl4 activity found by ion exchange chromatography in that specific acid. The per cent of the PFF's Cl4 activity in that organic acid was calculated as the total C14 activity in the specific organic acid peak divided by the total C14 activity eluted from the ion-exchange resin. The per cent recovery of C14 from the AGl-X8 resin during organic acid separation was 89% i 4% (N=17). 7. Total C14 activity contained in organic acids.-- The organic acids were eluted from tube 30 up to tube 160 of the organic acid separation. The per cent of total C14 activity put on the column which appeared in tubes 31 through 160 was multiplied by the total C14 activity in 67 14 the PFF to give total C activity as organic acids in each PFF. 14 8. Total C activity contained in the glucose fraction.--The amount of purified neutralized PFF (in- cubated with Glucostat) placed on the AGl—X8 resin for gradient elution of gluconic acid was considered to con— tain 5 m1 of neutralized PFF. Since some solution was probably left in the 5 purification resin columns used to remove organic and amino acids, the total glucose is underestimated even though Cl4—glucose was almost com— pletely recovered when passed one time, with a small water wash, through each resin. The C14 activity measured in the 3 ml of pooled eluted gluconic acid plated was corrected to the total 100 m1 volume collected from the resin column by multiplying by 100/3. Each Cl4 value so obtained was then calculated to represent the total C14 in the PFF by the ratio (total volume of PFF)/5 ml. Since the procedures used gave 85% recovery of C14- glucose and Cl4—gluconic acid from the purification resins and 79.0% conversion of Cl4—glucose to gluconic acid, the activities were finally multiplied by the cor- rection factor 1.50 to account for lack of 100% recovery and conversion. 9. Statistical analysis.——Significant difference between control and acidotic groups for renal glucose 68 production was tested with the Student's t Test (paired data). Statistical analysis of glycogen data was per— formed by Analysis of Variance. Standard errors (SE) were calculated with the following formula: SE = 2(x2) — where N is the number of observations N is less than 30 X is the individual observation. RESULTS I. Effect of Whole Body Acidosis on Renal Glucose Production Studies in the past using 1n 2122 kidneys have mea- sured renal glucose production during control periods and after the dog was made chronically acidotic by intra- gastric administration of ammonium chloride (Steiner 22 21., 1968; Churchill and Malvin, 1970a). The consensus of these 1n_2122 studies and of 1n 21222 studies is that chronic metabolic acidosis either reduces the renal uptake of glucose or increases its production. The data in Table 2 is from dogs in which renal glucose production was measured during normal acid-base balance and after making the dogs acutely acidotic by intravenous administration of HCl for one hour. This method of acidifying the blood is similar to the intra- gastric administration of ammonium chloride used in the studies of chronic acidosis (Steiner 22 21., 1968; Churchill and Malvin, 1970a), because in the dog the liver converts ammonium chloride to HCl and urea. A renal glu- cose mean uptake of 0.92 mg/min occurred during the con— trol period but one hour of acidosis was sufficient to cause a significant (P < .01) change to a renal glucose mean production of 0.68 mg/min. This result is similar 69 70 to that seen in previous studies with chronically acidotic dogs and the net renal glucose production found (Table 2), as expected from previous work (Steiner 22 21., 1968), was small. Although collateral circulation was eliminated when possible, lymph flow may not have been eliminated. If lymph flow from the kidney had occurred in increased amounts during acidosis, this could cause increased renal venous blood hemoconcentration and loss of glucose via the lymph. Haddy 22 21. (1958) showed that lymph flow increases as renal venous pressure increases. Lymph flows of 0.2 ml/min were measured during increased renal venous pressure (20 to 30 mm Hg). Even if lymph flows were 0.2 ml/min in the experiments in this thesis, this represents only 0.33% of the average renal blood flow of 60 ml/min. A renal venous hemoconcentration of 0.33% is less than the error encountered in the glucose analysis of a single paired blood sample and thus would not significantly af- fect the results in this thesis. Lymph flow was not mea- sured in the experiments presented here so its contribu— tion to renal venous hemoconcentration is not known. However, it is not probable that it could not account for the renal venous-arterial glucose concentrations measured which were as great as 12.5 mg % in some single V-A glu— cose concentration analyses. 71 Acute whole body acidosis did not affect the glycogen concentrations in the kidney (Table 3) indicating that glycogen did not contribute to the glucose release seen in the studies shown in Table 2. If only glycogen had con- tributed to the glucose formed (0.68 mg/min) for the hour of whole body acidosis, a total of 40.8 mg (0.68 mg/min) X (60 min) of glycogen would have to be released. The glyco- gen content of control dogs was 0.285 mg/g tissue. For a 56 g kidney (mean kidney weight for the dogs used in Table 2) the total pool of glycogen would be 16.0 mg which is not sufficient to account for the glucose release. Fur— thermore, the data shown in Table 3 indicate that acute acidosis did not significantly change the renal glycogen level. Likewise, in chronic metabolic acidosis it was found that total carbohydrate content (glycogen plus glu- cose) did not change significantly after 3 days of am— monium chloride acidosis (Steiner 22 21., 1968). The increased glucose observed to be released from the kidney during acute and chronic acidosis then must be newly formed within this organ. In addition to the renal metabolic effects des— cribed above, acute acidosis caused cardiovascular changes (Table 4). Six of the 7 dogs tested showed a decreased mean systemic blood pressure during the 2 hours of acidosis. Renal perfusion pressure, however, increased in 5 of the 8 dogs tested. Since blood flow was constant 72 and equal during both periods, the increased perfusion pressure could be attributed to increased renal resist— ance and/or blood viscosity. II. Effect of Local Renal Acidosis on Renal Glucose Production In contrast to the effect of whole body acidosis, one hour of local renal acidosis did not change renal glucose production significantly (Table 2). Also this group of dogs had a net glucose production of 0.32 mg/min during control conditions compared to the group used for whole body acidosis which had a net glucose uptake of 0.92 mg/min. The glycogen content of the kidneys after local acidosis was not significantly different from con- trols (Table 3). Local acidosis caused inconsistent cardiovascular effects in the systemic circuit (2 of 8 dogs had increased mean systemic pressure, 3 of 8 had decreased pressure, and 3 of 8 showed no change). Alterations in renal perfusion pressure and resistance were similar to the effects of whole body acidosis with 5 of 8 dogs having increased renal resistance during local acidosis (Table 4). Cl4—Glutamine Distribution and Metabolism in Whole Blood The purity of the Cl4—glutamine was first determined III . by amino acid chromatography. This purify check revealed that glutamine contained 87% of the Cl4 activity, organic 73 acids and neutral compounds 4.0%, and no activity was found in glutamic acid unless the labeled glutamine was allowed to remain unfrozen. The remaining 9.0% of the activity was located between the organic acid fraction and the amino acid fraction on the amino acid chromatogram. Although this labeled impurity was not identified, its elution pat- 14 peaks in the amino acid tern was known; and hence C chromatogram of the experimental blood filtrates could be attributed to its presence in the injection solution. Cl4—glutamine purity was claimed as 98% by New England Nuclear from their paper chromatography analysis. There- l4-glutamine fore, the results here indicate either (1) C is not stable in aqueous solution in which is was de— livered and/or (2) that the amino acid analysis employed here caused degradation of glutamine. The major impurity in the Cl4-glutamine identified by New England Nuclear was Cl4-pyrrolidone 5-carboxylic acid. This was confirmed by gradient elution anion-exchange chromatography. After Cl4—glutamine was incubated with whole blood for 5 minutes, the Cl4 activity still remained almost com- pletely in glutamine (Table 5). Less activity was present in glutamine (82.0% and 80.0% in whole blood compared to 87.0% in the purity check) and more activity in the I organic acid fraction and unknown fraction. The incuba— tion produced a small amount of glutamic acid in 74 Experiment B (0.5% of the total activity) but no other amino acids became labeled. The total activity in one ml of whole blood could be totally accounted for in the plasma of one m1 of blood (107% and 103% recovery of activity in plasma). From this data the proposed 1n 2122 renal metabolism experiments ap- 14 peared feasible because (1) the injected C -glutamine would remain in the plasma and be available for renal up— take and (2) there was little formation of Cl4-glutamic acid or other degradation products after 5 minutes of incubation in plasma. The amount of Cl4—glutamic acid present is important because the kidney can have a net uptake of glutamic acid under certain conditions (Shalhoub g 21., 1963) . IV. Study of Cl4-Glutamine Metabolism in the In Situ Pump—Perfused Dog Kidney Cl4-glutamine metabolism by the 12 situ pump- perfused dog kidney was studied in 3 experiments. Ex- periments I and III were performed on dogs at normal blood pH (7.35 and 7.35) and Experiment 11 on a dog made acutely acidotic (blood pH 7.18) by intravenous infusion of hydro- chloric acid. No statistical comparisons between control dogs and the acidotic dog will be made because of the nature of the data and the small number of dogs used. 14 Using a pump-perfused kidney most of the C label injected can be recovered. Tables 6, 7, and 8 show that 75 99.2%, 97.8%, and 104% of the total C14 label injected was recovered either as protein-free filtrate or as precipitate from the blood—perchloric acid mixture. Loss of C1402 was not measured in these experiments and loss due to experimental technique is estimated at approximately 1% of the total tissue collected. Blood which drained from the kidney (as it was being rapidly extracted from the body) would account for some Cl4 loss. Most of the Cl4 activity was present in the protein—free filtrate (Experi— ment I-93%, II-91%, III-97%) in which C14-labe1ed amino acids, organic acids, and glucose were identified. The C14 activity in the precipitate may be due to (1) labeled compounds which are insoluble in the acidic filtrate, (2) trapped filtrate within the precipitate, and (3) filtrate contamination because of incomplete decantation. 14 activity were mea- Three distinct fractions of C sured in the protein-free filtrates: amino acids, organic acids, andflglucose (Tables 9, 10, and 11). In each experi— ment the amino acid and organic acid fraction had high activity in the first blood sample and decreasing activi— ties in the last 4 samples. This probably is due to the rapid passage of the injection solution through the renal cortical vasculature followed by the injection solution's slow passage through the renal medullary vasculation dur— ing the later sampling. 76 Cl4—g1ucose, on the other hand, showed the reverse pattern where the first blood sample contained little Cl4-glucose and the latter samples contained increasing amounts of Cl4—glucose. This is especially apparent in 14 Experiment I where C activity in glucose increased 20— 14 fold between blood sample 1 and 5. That the C activity in glucose resulted from renal glucose production is indi- cated for the following reasons: (1) Although Cl4-glucose was not specifically analyzed with Glucostat in the 12 l4-glutamine with whole blood, the Vitro incubation of C glucose fraction was isolated from amino acids on the amino acid column. No C14 activity was present in either of the 2 experiments indicating no Cl4-glucose production by whole blood. (2) When injection solution was purified through the 5 resin beds, only 0.15% of the Cl4 activity was left. 14 This would be the maximum amount of C —glucose activity which could be present in the injection solution. The total glucose fraction in Experiments 1, II, and III was equal to 3.31%, 0.43%, and 0.63% of the injected Cl4 activity. In all cases, and especially in Experiment I, more glucose was measured than could be accounted for by a Cl4—glucose impurity in the injection solution. (3) Finally, if this fraction had been an impurity in the injection solution, one would expect it to pass through 14 the kidney in a pattern similar to C -g1utamine or pyrrolidone 5—carboxylic acid (an impurity formed .I '#I 77 spontaneously from glutamine) with most appearing in the first blood sample. Instead, the smallest amount of glu— cose is found in the first blood sample and the most in the latter samples. The data then indicate production of C14— glucose from extracted Cl4-glutamine and release of this glucose into renal venous blood. However, during Experi- ment I and II there was no net production of glucose by the kidney when measured by V—A glucose analysis (Table 19). The kidney extracted 29.8%, 39.0%, and 24.0% of 14 the injected C -glutamine in Experiments 1, II, and III. Kidney tissue and renal venous blood were analyzed for Cl4-labeled amino acids (Tables 12, 13, and 14) and C14- 1abeled organic acids (Tables 15, 16, and 17) to determine the fate of the Cl4 extracted. The amino acid fraction contained only C14 activity in glutamine, glutamic acid, and aspartic acid. C14 in specific amino acids was de- tectable at the level of approximately 0.05% of the activ- ity injected. Only in Experiment II was there more C14- glutamic acid in renal venous blood than was present in the injection solution (Table 13). In the kidney tissue 14 activity appeared in glutamic acid. The significant C high Cl4-glutamic acid activity in the injection solution of Experiments I and III is unexplained since it occurred neither in Experiment II nor in the 1n Vitro whole blood Experiments A and B. All procedures with respect to 78 filtrate preparation and amino acid chromatography were identical in all experiments. However, it has been shown that glutamine can spontaneously break down producing am— monia even at temperatures near 0° C which may explain at least part of the Cl4 —glutamic acid production (O'Donovan and Warner, 1969). Aspartic acid, which could not clearly be separated on the amino acid chromatogram because of its close geo— metric proximity to glutamine and because of glutamine's overwhelmingly high activity, was separated by organic acid chromatography. No Cl4—aspartic acid was present either in the injection solutions or in any renal venous blood samples. However, Cl4-aspartic acid was identified in the kidney tissue. When the neutralized PFF's of renal venous blood samples were chromatographed on AGl-X9 resin, there were no definitive peaks of Cl4 activity in the organic acid fraction except for pyrrolidone 5—carboxylic acid and an unidentified impurity. This was also true for the organic acid chromatograms of the injection solutions. However, the injection solutions and the neutralized PFF's of renal 14 activity in the organic acid venous bloods did contain C fraction when the total C14 activity across the entire fraction was determined (Tables 9, 10, and 11; Methods Section III, C, 8). As a result, the organic acid frac— tion was divided into areas containing known organic 79 acids (determined from carrier organic acid analysis) and the Cl4 activity determined in each area (Tables 15, 16, and 17). Each individual area, which is denoted by the major organic acid contained in that area, may contain other compounds in addition to the organic acid for which the area was named. For example, the area denoted as d— KG could contain d-KG as well as other unknown substances eluted from the resin in the same position as a-KG. The delineation of these areas was based on the elution pat— tern of carrier organic acids which had been added to the neutralized PFF's. The renal venous blood data show that the malate, citrate and/or pyruvate, and fumarate areas consistently contain more C14 activity than was present in these areas of the injection solution chromatogram for Experiments I and 11 (Tables 15 and 16). In Experiment III there was more C14 activity in the succinate and citrate areas, than was present in the injection solution (Table 17). This data suggests release into the renal venous blood of C14- 14 14 malate, Cl4—pyruvate and/or C —citrate, and C —fumarate produced by the kidney. However, there also is evidence which refutes this data: (1) There were no definitive Cl4 activity peaks in any of the neutralized PFF's of renal venous bloods chromatographed. This suggests that the Cl4 activity in each area is not concentrated in a particular compound but rather in several compounds or 80 in a compound which is not eluted as a distinct peak. (2) In general, the Cl4 activity in each area was highest in the first blood sample collected with a progressive decrease in the subsequently collected blood samples. This is the pattern expected for a compound which passes through the kidney unaltered. (3) If the Cl4 activity for all 5 blood samples is summed for each area and expressed as a per cent of the C14 activity present in the injection solution for that area (Table 18), there are no areas (except the unknown and fumarate areas) which are notably different from 100%. A value of 100% would represent no uptake or production of the substance injected. There- fore, it cannot be definitely stated that Cl4-labeled organic acids are released into the renal venous blood by the kidney with the possible exception of fumarate and citrate. However, in the kidney tissue there was definite Cl4 activity in pyrrolidone 5—carboxylic acid, the unknown compound, succinate, malate, citrate and/or pyruvate. The C14 activity in pyrrolidone and the unknown can be ac— counted for by their presence in the injection solution. In most cases there was more C14 activity in tissue suc— cinate, malate, citrate and/or pyruvate than was present in the injection solution (Tables 15, 16, and 17) suggest- l4 ing C —labe1ing of these organic acids in the kidney. 82 TABLE 2.——Cumulative Data from the Experiments on the Ef— fect of Whole Body and Local Renal Acidosis on Renal Glu- cose Production Experiment Control2 Acidosis2 P Whole Body Acidosis (8)l Blood pH Systemic 7.38-7.53 7.08—7.33 Renal Arterial 7.38-7.53 7.08—7.33 Renal Blood Flow (m1/min)359:2 59:2 NS Renal Glucose Production ug/min/g tissue4 —15.8 +2.8 14.5 i5.o .01 mg/min — 0.92¥0.18 0.68+0.23 .01 Urine Flow (ml/min) 0 0 Local Renal Acidosis (8)l Blood pH Systemic 7.40—7.59 7.38-7.51 Renal Arterial 7.40-7.59 7.13-7.30 Renal Blood Flow (ml/min)3 57+l 57:1 NS Renal Glucose Production _ ug/min/g tissue4 5.2 +7.5 5.1 :6.6 NS mg/min 0.32E0.38 0.31:0.32 NS Urine Flow (ml/min) 0 0 1Number of animals. 2Values are 2 SE except for pH's which are given with their ranges. 3Renal blood flow was constant for each dog during control and experimental periods. 4A positive value indicates glucose production and a negative value indicates glucose uptake. In both groups mean renal glucose production was calculated for each individual dog. 83 TABLE 3.--Glycogen Content of Kidney in Control, Whole Body Acidosis, and Local Acidosis Glycogen Contentl Dog # (Mg/g tissue) Mean : SE Control Period A left kidney 129 B left kidney 99 B right kidney 80 C left kidney 320 C right kidney 276 D left kidney 728 D right kidney 366 285 i 85 Whole Body Acidosis3 6 223 8 476 9 186 2 11 280 291 i 64 Local Renal Acidosis3 3 340 4 76 5 250 6 272 7 393 2 11 193 254 + 45 lGlycogen values are those measured X 2.3 to correct for less than 100% extraction of glycogen. 2Values are not significantly different from con- trols. P > .10. 3 All kidneys used were the left experimental kidneys. 84 TABLE 4.——Effect of Whole Body and Local Acidosis on Mean Systemic and Renal Perfusion Blood Pressure Mean Renal Renal Systemic Perfusion t H3 (mm Hg) (mm Hg) ar ‘ 9 Whole Body Acidosis # 1 c1 — 60 7.42 A2 — 60 7.14 # 3 C 150 75 7.46 A 125 120 7.22 # 4 C 60 130 7.46 A 125 120 7.19 # 6 C 160 75 7.46 A 100 125 7.26 # 7 C 120 60 7.38 A 100 60 7.26 # 8 C 150 75 7.53 A 60 210 7.33 # 9 C 150 60 7.50 A 110 80 7.31 #11 C 110 50 7.42 A 50 75 7.08 Local Renal Acidosis 3 C 125 45 7.42 A 75 120 7.13 # 4 C 60 45 7.40 A 70 70 7.30 # 5 C 130 45 7.46 A 100 45 7.13 # 6 C 125 40 7.59 A 125 50 7.20 # 7 C 125 65 7.45 A 105 100 7.20 # 8 C 85 45 7.49 A 50 35 7.19 #10 C 120 105 7.49 A 120 90 7.22 #11 C 145 65 7.49 A 145 75 7.18 lControl period. 2Acidosis period. 3 Taken before blood entered Sigma pump. During local studies renal pH was taken directly before entrance into kidney (these are listed under A). Whole body pH did not change more than i .05 during local acidosis. 85 TABLE 5.—-Cl4—Glutamine Distribution and Metabolism in Whole Blood Experiment A Experiment B Counts/minute in 1 ml plasma 1,364,500 1,424,100 Counts/minute in 1 ml whole blood 912,200 919,300 Counts/minute expected in the plasma of 1 m1 whole blood 982,400 947,026 Final hematocritl 28.0% 33.5% Per cent recovery of counts in plasma 107.9% 103.0% C14 amino acid analysis of whole blood2 Glutamine 82.0% 80.0% Glutamic Acid 0.0% 0.5% Organic Acid Fraction 7.0% 9.0% Glucose Fraction 0.0% 0.0% Unknown Peak 9.0% 10.5% 1Refers to the hematocrit measured after the iso— tope had been added to the 7 ml of whole blood. 2Per cents given are based in the total counts in each fraction divided by the total counts in whole blood. 86 TABLE 6.-—Total Cl4 Activity in Protein—Free Filtrates and Precipitates from Experiment I Total C14 Activity (counts/minute) Protein—Free Filtrate PreCipitate Blood-1 min 24,882,498 1,019,271 Blood—2 min 6,707,136 284,427 Blood—3 min 2,500,664 91,080 Blood—4 min 1,598,113 44,583 Blood-5 min 1,695,842 67,188 Kidney Tissue 9,510,739 1,429,643 Total Activity Injected 50,200,785 counts/minute Total Activity Recovered 49,795,184 counts/minute Per Cent Activity Recovered 99.2% 87 TABLE 7.——Tota1 Cl4 Activity in Protein-Free Filtrates and Precipitates from Experiment II Total C14 Activity (counts/minute) Protein-Free Filtrate Prec1pitate Blood-1 min 15,919,982 873,906 Blood-2 min 11,719,858 715,028 Blood—3 min 5,030,243 201,960 Blood-4 min 3,113,208 165,924 Blood-5 min 3,338,481 119,543 Kidney Tissue 13,803,833 2,130,084 Total Activity Injected 58,404,159 counts/minute Total Activity Recovered 57,132,050 counts/minute Per Cent Activity Recovered 97.8% 88 TABLE 8.—-Tota1 Cl4 Activity in Protein—Free Filtrates and Precipitates from Experiment III Total C14 Activity (counts/minute) Protein—Free Filtrate PreCipitate Blood—l min 26,447,925 2,705,637 Blood-2 min 7,398,677 448,800 Blood—3 min 2,316,064 232,452 Blood—4 min 1,495,669 76,280 Blood—5 min 748,836 38,115 Kidney Tissue 6,466,851 345,345 Total Activity Injected 46,556,443 counts/minute Total Activity Recovered 48,720,651 counts/minute Per Cent Activity Recovered 104% _.,.- 1.--- .1_.5?i}: 89 TABLE 9.--Cl4 Activity of the Amino Acid, Organic Acid, and Glucose Fractions in the Blood and Kidney Samples from Experiment 1 . Amino Acid Organic Acid Glucose Filtrate Fraction Fractionl Fraction1 Blood—1 min 19,806,468 2,662,427 21,494 79.6 10.7 0.9 Blood—2 min 5,204,738 462,792 282,029 77.6 6.9 4.2 Blood—3 min 1,472,891 237,563 368,678 58.9 9.5 14.7 Blood—4 min 730,338 46,345 400,398 45.7 2.9 25.1 Blood—5 min 469,735 214,120 438,384 28.3 12.9 26.4 Kidney Tissue 7,009,415 1,483,675 149,163 73.7 15.6 1.6 1The top number is the total C 14 minute) in the filtrate as this fraction. her is the per cent of total C14 activity in the filtrate as this fraction. activity (counts/ The lower num— 90 TABLE 10.-—Cl4 Activity of the Amino Acid, Organic Acid, and Glucose Fractions in the Blood and Kidney Samples from Experiment 11 Filtrate Amino Acid Organic Acid Glucose Fraction Fraction Fraction Blood-1 min 11,605,667 2,817,837 8,603 72.9 17.7 0.5 Blood-2 min 8,778,174 2,929,965 32,841 74.9 25.0 2.8 Blood-3 min 3,968,862 1,091,563 24,840 78.9 21.7 4.8 Blood—4 min 2,465,661 703,585 65,240 79.2 22.6 2.0 Blood—5 min 2,717,524 383,925 42,558 11.5 1.3 Kidney Tissue 11,581,416 2,194,809 76,740 83.9 15.9 5.5 1The top number is the total C 14 minute) in the filtrate as this fraction. her is the per cent of total C14 activity in the filtrate as this fraction. activity (counts/ The lower num— 91 TABLE ll.-—Cl4 Activity of the Amino Acid, Organic Acid, and Glucose Fractions in the Blood and Kidney Samples from Experiment III Filtrate Amino Acid Organic Acid Glucose Fractionl Fractionl Fractionl Blood—1 min 22,057,569 1,957,146 21,042 83.4 7.4 0.8 Blood—2 min 6,370,261 547,502 46,242 86.1 7.4 0.6 Blood-3 min 1,880,644 252,451 66,528 81.2 10.9 5.1 Blood-4 min 1,154,656 116,662 70,728 77.2 7.8 4.8 Blood-5 min 556,385 98,098 47,564 74.3 13.1 6.4 Kidney Tissue 5,483,890 982,961 38,100 84.8 15.2 5.9 1The top number is the total C14 activity (counts/ minute) in the filtrate as this fraction. The lower num- ber is the per cent of total C14 activity in the filtrate as this fraction. 92 TABLE 12.——Cl4 Activity Contained in Each Amino Acid in the Blood and Kidney Samples and Injection Solution from Experiment I Amino Acid Filtrate . A t' Glutaminel Glutamatel Spa? 10 Ac1d Blood—1 min 18,164,223 671,827 —— 73.0 2.7 Blood—2 min 3,950,503 241,457 —- 58.9 3.6 Blood—3 min 1,107,794 72,519 —- 2.9 Blood—4 min 476,238 —- -- 29.8 -- Blood—5 min 448,157 53,115 —- 27.0 3.2 Kidney Tissue 2,025,787 4,346,408 133,150 21.3 45.7 1.4 Injection Solution 34,387,538 5,412,685 —— 68.5 10.8 1The top number is the total C14 activity (counts/ minute) in the filtrate. The lower number is the per cent of total C14 activity in the filtrate as this fraction. 93 TABLE 13.—-Cl4 Activity Contained in Each Amino Acid in the Blood and Kidney Samples and Injection Solution from Experiment II Amino Acid Filtrate Glutaminel Glutamate1 Aspartic Ac1d Blood—1 min 10,236,548 334,320 -- 64.3 2.1 Blood—2 min 7,852,305 292,996 -- 67.0 2.5 Blood-3 min 4,139,890 241,452 —— 82.3 4.8 Blood—4 min 2,478,114 136,981 —- 79.6 4.4 Blood-5 min 2,757,585 166,924 -— 82.6 5.0 Kidney Tissue 6,391,175 4,872,753 220,861 46.3 35.3 1.6 Injection Solution 45,029,606 642,446 -— 77.1 1.1 1The top number is the total C14 activity (counts/ minute) in the filtrate. The lower number is the per cent of total C14 activity in the filtrate as this fraction. 94 TABLE 14.-—Cl4 Activity Contained in Each Amino Acid in the Blood and Kidney Samples and Injection Solution from Experiment III Amino Acid Filtrate A t' Glutaminel Glutamatel spar 1C Ac1d Blood-1 min 19,915,288 899,229 -- 3.4 Blood-2 min 5,807,961 310,744 —- 78.5 4.2 Blood—3 min 1,579,556 60,218 —- 68.2 2.6 Blood—4 min 1,014,064 ‘82,262 —— 67.8 5.5 Blood-5 min 560,129 24,712 —- 74.8 3.3 Kidney Tissue 2,884,216 2,360,400 87,069 44.6 36.5 1.3 Injection Solution 37,990,057 3,165,838 '- 81.6 6.8 1The top number is the total C14 activity (counts/ minute) in the filtrate. The lower number is the per cent of total C14 activity in the filtrate as this fraction. .QOAuDHom soauowflch .sOfluOMHm many mm mumsuHHw map :H mpfi>fiuom «Ho HMHOp mo ucwo Mom was we HmnEds Hw3oa GAB .wumuuaflm may CH AwBSCHE\mucsoov mufl>fluom o Hmuou 6;» ma Hones: mow wcfia 95 va o>.o v0.0 mH.o om.o mo.m mv.o om.oa mm.o mov.amm omo.om me.mm How.ooa Hmm.mvm.a vom.mmm oma.mwm.m mmm.amm «.m.H mm.o mw.o mv.m mo.m MH.H wm.m mN.H on.o mmw.hw mam.mm mmm.wfim mmo.amm Hhv.hoa mom.wmv www.maa mnm.mw mammfle mmcpflm nm.m Hm.H In vm.o mv.H wH.N hm.m Hm.m wmw.mm mmn.om I: mow.mH onm.mm Hmm.mm hum.vm Nvm.mm QHE m cooam II II wv.m II II I: 05.0 mm.o I: ll vao.mm :I II I: wwH.HH vaa.m CHE v poon on.m mm.H m>.H oo.H om.m mm.o on.m ma.o mmm4Nm www.mv Hmm.mv moo.mm omm.on mmn.m mmm.mwa ooo.m QHE m pooam ww.o mw.o wb.o II «v.0 ba.o om.v hm.o mmm.om mmw.mm onm.om II Ham.mm Nov.HH mvm.amm omm.mm CHE N poon mm.o mH.o av.o mv.o on.H vm.o mm.n wm.o moo.hm NHw.mm wao.moa HFm.HHH moo.mmv oom.vw www.mmw.fi mHo.mom see a poon 6 pflo< H um>DH>m m I OM15 wuwumfism a m¢muuao Hmumamz HEBOGMQD Hmumcfloosm UAH xonnmo m wamuomq H H . HGGOpflaonhmm mgmupaflm me< cflod oasmmno H meEflHmmxm EOHM coauSHom QOHuowncH UGM memEMm wmnpflm paw pooam wag CH me< ©H0< UqumHO Somm cH U®CkuGOU hpfl>flpo< «HUII.mH mqmdfi 96 .COHusHom COHpooflCHN .COHuomum mHCw mm >CH>Hvom 0H0 Hmnou mo quo Com mCu mH umCECC CmBOH mCB .wumubHHm mCu CH AmCSCHE\muCCOUV huH>Huom o ngou wag mH HwQECC mow 6C9 VH H hm.o vm.o NN.0 Hm.o Hm.m 0m.m 0N.MH mm.o mHH.mom onH.00H mmv.me Hom.>0m bmH.Hv0.H mmm.m>m.H mvm.moh.n N0>.m0m N.m.H v0.0 N0.0 00.N 0N.m Hm.0 0H.v 0>.m 0H.o www.mNH wwm.mw 000.0mm mmn.va me.mmH mnv.Hhm 000.00m mvw.vm mummHB thUHx mm.m H0.H mm.0 00.0 mv.0 mh.0 hv.m NN.0 0MH.vm mHn.mm www.mH Hm0.mm NN0.0H mmo.mm mH0.NmH 00m.» CHE m woon mw.H 0N.o mm.H wv.H 00.N 0m.0 0H.mH mm.o vmm.»m «00.0 www.mm mno.0v NMH.vm www.mH «00.0nw mmH.0H CHE q poon Hm.0 w>.o Hw.0 00.0 mm.H hm.o 0m.nH 0H.H 000.0m mmm.mm vmo.om mon.vm 050.05 NHO.mH mmm.0hm Nmm.mm CHE m toon 00.0 vm.0 05.0 >0.H vv.m no.0 0m.nH 00.0 n0H.0HH wNH.mN www.mm Nov.mNH m0H.mov m00.MHH mnm.0m0.m 0N0.m0H CHE N tOOHm hm.0 mm.0 «v.0 00.0 00.m 00.0 om.HH 00.0 www.00 0H0.0m «No.0HH 000.HOH www.mmv 0mm.wOH wnv.vmm.H Hmm.mmH CHE H poon 6 ©Ho< um>5Hmm oxla mpmumECm H 0 mm H quHmz CSOCxCD wmeHoosm UHmeonumolm mumuomq H H a p u.u H H H HmCOUHHOHumm H mpmsuHHm mmufi pHofi OHCM0HO HH quEHHmmxm Eoum COHuCHom COHuommCH 0C8 meQEMm thpHM UCM poon 0:» CH mmum UHom UHCmmHO Comm CH prHmuCOU wuH>Huo¢ «H011.0H mHmCB 97 .COHudHom COHuowflCH N .CoHuomuw mHCu mm >HH>Huom 0H0 Hmuou wo quo Hmm wCu mH HwCECC Hm30H mCB .mpmnuHHw mCu CH AmusCHE\muCCoov >UH>Huom o Hmuou 0C“ wH HmCECC mow mCB 0H H H0.0 00.0 00.0 00.0 00.H 0N.0 HH.N 00.0 000.00N 000.00H 000.N0 000.00N 000.000 000.00H H00.N00 000.00N N.m.H 00.H 00.0 00.0 00.N 00.0 00.H 00.N H0.0 000.00 0N0.N0 000.0HN 0H0.NOH 0NN.00N 00N.00H 000.00H 000.00 mammHB >wC©HM 00.N 00.H 0N.0 II 00.N In 00.0 00.0 000.0H 00N.0H 0H0.N II NOH.HN II 000.00 000 CHE 0 GOOHm 00.N HH.H 00.0 00.0 II 00.0 00.N 00.0 000.H0 N00.0H 000.0 000.HH II 0HN.HH 000.00 000.0 CHE 0 UOOHm 00.N H0.0 00.0 00.0 I: 0H.0 00.0 0N.0 00H.00 000.HN 000.0H H00.0 || 00H.0 H00.00H N00.0 CHE 0 UOOHm 00.0 00.0 II 0H.0 00.0 0H.0 00.0 0N.0 0H0.00 0H0.H0 II 000.HH 000.00 000.0H 00H.N00 000.0H CHE N UOOHm H0.0 0N.0 N0.0 00.0 00.H 00.0 0N.0 00.0 000.H0 000.00 000.00 N00.00H NON.0H0 000.0NN 0H0.0HH.H NHN.00 CHE H Uoon CHUC Humid HmHMHmfidh MQWMMMMMW mUMHmz C3OCxCD mCMCHousm UHH>XOAHMUI0 wpmuomq . H H H HmCowHHounhm H mumupHHm mmhm UHU¢ UHCmmuo HHH quEHHmmxm Eouw COHuSHOm COHprmCH 6C0 memEmm mepHM UCM woon 0C0 CH mmum 0H0< UHCmmuo Comm CH meHMCCOU muH>HCom UII.0H mqmde 0H 98 TABLE 18.——Tota1 Cl4 Activity in Renal Venous Blood Samples 1 Through 5 in Each Organic Acid Area Expressed as a Per Cent of the C 4 Activity in the Injection Solution for That Area orgagigaACid Experiment Mean : SE I II III Lactate 88.5 110.0 50.4 83.0 + 17.3 Pyrrolidone 5-Carboxy1ic Acid 28.4 70.8 173.0 90.7 + 41.9 Succinate 62.2 20.4 234.0 105.5 + 65.2 Unknown 41.0 59.8 57.7 52.8 + 5.1 Malate 152.0 128.0 65.8 115.2 + 25.6 Citrate 264.0 218.0 340.0 274.0 + 35.6 Fumarate 725.0 103.0 97.0 308.0 + 206.0 a—KG 61.8 73.4 94.0 76.4 + 24.0 TABLE 19.—-Genera1 Data for Experiments I, II, and III on of Cl4—G1utamine Exp. Exp. Exp. I II III Dog Weight (lbs) 50 50 42 Dog Sex F M F Renal Blood Flow (ml/min) 62 60 60 Blood pH 7.36 7.18 7.35 Kidney Weight (g) 83 109 97 Glutamine Concentration 0.38 0.39 0.40 Injection Solution (UM/ml) Respiration Rate (breaths/min) 25 14 20 Tidal Volume (m1) 400 300 350 Systemic Pressure2 125 120 100 Renal Arterial Pressure2 90 75 65 (mm Hg) Volume of Injection Solution 2.25 2.36 2.05 given (m1) Renal Glucose Production 0 O - (mg/min) lpH measured within 5 minutes before injection solu- tion given. 2Pressure within 5 minutes before injection solution given. DISCUSSION I. Effect of Whole Body and Local Renal Acidosis on Renal Glucose Production This group of experiments was designed to determine whether acute whole body and/or local renal acidosis would affect renal glucose production and to compare the effect to chronic (3 days) metabolic acidosis. Steiner et a1. (1968) have shown that chronic metabolic acidosis increases glucose production from the kidney, although the values obtained are barely significant (P < .025). Roxe 33 21. (1970) and Churchill and Malvin (1970a) found no change in glucose production during chronic metabolic acidosis. The acute effects of whole body acidosis described in this thesis support the findings of Steiner 23 31. (1968) in the in yizg kidney and are in agreement with the find- ings of Goorno et 31. (1967) and Churchill and Malvin (1970a) that chronic acidosis stimulates glucose produc- tion in dog renal cortex slices. Considering all the studies available for the dog, it appears that both chronic and acute metabolic acidosis cause either small increases in net renal glucose production or small reduc— tions in net uptake. When only the renal arterial blood was made acidotic, no change in renal glucose production occurred (Table 2). 100 101 There are no other in 2129 studies of local renal acidosis with which to compare these data. The cause of the whole body acidosis effect remains uncertain but the lack of a local kidney acidotic effect suggests that renal acidosis per se is not the direct cause. This then implies either an effect of innervation (no information is available on the role of renal nerves in renal glucose production) or blood—born metabolites and hormones. Acidosis may exert its effect elsewhere than the kidney and the kidney is indirectly affected by one or all of these factors. To show the effects of a hormone or metabolite it would be necessary to measure its blood level or turnover concomit— antly with changes in renal glucose production. In the experiments presented in this thesis there is evidence that the last paired blood sample drawn 2 hours after the initiation of local renal acidosis had a signifi- cantly greater V—A glucose difference than the first 3 samples drawn (data not presented). If the study had been extended for longer than 2 hours, an increased glucose production by the kidney may have been seen. This again suggests that a systemic factor is reinforcing the effect of acidosis on the kidney. It is also possible that blood glutamine levels were depleted during the first 2 hours of local renal acidosis. The liver is stimulated to increase production of glutamine during metabolic acidosis but the precise mechanism is not known. In the local acidosis 102 studies there may not have been a stimulation for in— creased hepatic glutamine synthesis. Addae and Lotspeich (1968) suggest that the increased blood ammonia levels from the kidneys, brain, and gastrointestinal tract stimu— late hepatic glutamine production. In the local acidosis studies only the experimental kidney would increase am— monia production. Thus, the blood ammonia levels may not rise as much as when both kidneys, the brain, and the gastrointestinal tract increase their ammonia production during whole body acidosis. As a result, the liver has no stimulus to increase its glutamine production. The fall in blood glutamine levels may then effectively re— move substrate for renal gluconeogenesis. Although glycogen concentrations were not affected by either whole body acidosis or local renal acidosis, the role of glycogen in renal glucose production (if any) is not yet certain. Glycogen levels may have changed transiently during the acidosis but returned to normal by the end of the experiment. If renal glucose production is the result of increased renal utilization of glutamine or some other substrate which is converted to glucose, then glycogen may not be involved. However, the newly formed glucose may still be involved in the constant production and breakdown of renal glycogen. During whole body acidosis significant decreases in the mean systemic blood pressure occurred while there was 103 generally an increase in renal perfusion pressure and renal vascular resistance (Table 4). The pattern of re- duced systemic pressure and increased renal resistance was also seen by Bersentes and Simmons (1967) during metabolic acidosis. Renal responses to hydrogen ion are complicated by increased barostatic reflexes resulting from decreased systemic pressures and increased cate— cholamine secretion due to acidosis (Morris and Millar, 1962). Thus, any local effect of the hydrogen ion during whole body acidosis may be overshadowed by systemic ef- fects. When only the renal arterial blood was acidified, there was no consistent systemic effect, but in 5 of 8 dogs there was an increase in renal resistances. Again this effect may not be due solely to the vasoactivity of the hydrogen ion. When the local acidosis experiments were first tried, acid was infused directly into the per- fusion tubing causing immediate and massive increases in renal perfusion pressure. The cause is unknown but may have been either cellular breakdown releasing a con— strictor element, a physical blockage due to cellular destruction, or a blood reaction to the hydrogen ion. To overcome this difficulty, a mixing flask was placed in the circuit whereby acid could be added to a large volume of blood (100 m1) before being pumped to the kidney. Instead of completely eliminating the problem, the effect 104 might have been "diluted" but still causing some renal resistance increases. In both designs, with or without the mixing chamber, the renal arterial pH was the same suggesting that the renal vasoconstriction might have been caused by something other than local renal pH. II. Cl4-Glutamine Distribution and Metabolism in Whole Blood 14 A study of C —glutamine distribution and metabolism in whole blood was necessary to determine whether the pro— posed 13 vivo experiment was feasible. The 2 whole blood 1n vitro experiments showed that all the Cl4 activity in whole blood could be accounted for by the Cl4 activity in plasma (Table 5). Since all exogenous glutamine remained in the plasma after 5 minutes of incubation with whole blood, it is apparent that none entered erythrocytes and 14 that all C -glutamine injected would be available for extraction. Although Cl4—glutamine did break down slightly in whole blood, very little glutamic acid and no glucose was formed. Glutamic acid is important to the renal metab— olism study because the kidney can produce glutamic acid during normal acid—base balance and extracts it during acidosis (Shalhoub at 31., 1963). If Cl4-g1utamine had formed Cl4-glutamic acid in the blood, it would not be possible to distinguish this glutamic acid from that re— leased by the kidney arising from extracted Cl4—glutamine. 105 Even though the kidney has net production of glutamic acid, there may be uptake at one renal site and production 1 at another. The object of the C 4—glutamine study was to determine the products of its metabolism after renal ex— traction and not the products of extracted Cl4—g1utamic acid. Although a specific enzymic Cl4-g1ucose assay was not done in these 2 experiments, the area in which glucose separates on the amino acid chromatogram did not contain Cl4 activity. Any Cl4—glucose found in the 13 3129 ex— periments can therefore be attributed to renal metabolism. O'Donovan and Warner (1969) have shown that aqueous solutions of glutamine can form ammonia non-enzymatically. These data and the results found in these 2 experiments that 0.5% glutamic acid could be formed in dog whole blood 14 alone indicated that all C -g1utamine solutions injected into the dog should be chromatogramed. This would then act as an internal standard testing the amount of C14- glutamine breakdown occurring due to the total experimental and analytical techniques. III. Study of Cl4—G1utamine Metabolism in the l£.§i32 Pump‘Perfused Dog Kidney The primary aim of this group of experiments was to study the metabolic fate of Cl4-glutamine injected into the renal artery of the dog. It has been shown that glutamine is extracted by the kidney during normal acid-base balance and that extraction increases when an acute metabolic 106 acidosis is established (Addae and Lotspeich, 1968). It also has been amply demonstrated in dog renal cortex slices that glutamine can produce gluconeogenesis (Goorno gt g1., 1967; Churchill and Malvin, 1970a). A recent finding (Roxe gt g1., 1970) has indicated that C14- glutamine infusion into the renal artery of the dog is l4-glucose into the renal associated with the release of C venous blood within one minute after injection. Despite glutamine's known involvement in renal ammoniagenesis and gluconeogenesis (Steiner gt g1., 1968; Literature Review, Section III) the metabolic fate of its carbon skeleton has not been studied in the kidney. Preparation of an ideal physiological kidney for renal metabolic studies although desirable is highly im— practical. The pump-perfused 1g g1tg dOg kidney was the best choice for several reasons: (1) An 1g_g1tg prepara- tion is exposed to renal innervation and blood metabolites and hormones. (2) Injections can be made into the renal artery without affecting renal blood flow. (c) Cannula— tion of the renal vein and freezing the kidney after the experiment allows nearly total collection of the isotope injected. (4) The amount of surgery necessary to prepare the 1g g1tg kidney is comparable to the preparation of an isolated perfused rat kidney. The rat kidney preparation can be done without affecting the functional integrity of the kidney (Nishiitsutsuji-Uwo gt g1., 1967; Bowman, 1970). 107 (5) Recirculation of isotope and labeled products of renal metabolism is prevented. (6) The renal blood flow can be precisely controlled and measured without the use of PAH infusion. A large amount of isotope (250 no) could be injected quickly in a small volume (< 2.5 ml) directly into the renal artery to produce substantial labeling in the kidney. However, neither the degree of degradation of the extracted Cl4—g1utamine could be estimated nor could the amount of C14 activity released by the kidney into the renal venous blood be predicted prior to these studies. As a result of these experiments, several facts have been learned about renal Cl4—glutamine metabolism and improvements in the design can be suggested to study renal Cl4-glutamine metabolism more thoroughly. l4—glutamine was injected into When a solution of C the renal artery, the kidney extracted 29.8%, 39.0%, and 24.0% of the glutamine in Experiments I, II, and III. The acutely acidotic dog (Experiment II) extracted the greatest per cent of glutamine as would be expected from previous studies (Shalhoub gt g1., 1963) with chronic acidosis. The majority of the remaining Cl4 activity passed rapidly through the kidney in the first 2 minutes of blood collection (Tables 6, 7, and 8). The last 3 blood samples contained progressively less Cl4 activity showing that no major contribution to total blood Cl4 108 activity was being made by the kidney. This point is further substantiated by the Cl4 activity and content of the amino acid and organic acid fraction of the 5 one- minute blood samples. In the renal venous blood samples only the amino acids glutamine and glutamic acid contained Cl4 activity above 0.10% of the Cl4 activity injected (Tables 12, 13, 14). The C14 activity in these 2 amino acids is probably due entirely to their presence in the injection solution 14 because (1) their decreasing C activity from blood sample 1 to 5 follows the same pattern that is expected if no renal production occurs and (2) their total C14 activity in bloods 1 through 5 can be accounted for by their C14 activity in the injection solutions except for glutamic acid in Experiment II. From this data it ap- l4-g1utamine are pears that amino acids labeled from C not released in large quantities into the renal venous blood in the time course of these experiments. In some of the renal venous blood samples there was Cl4 activity in several of the organic acids as- sayed, but it cannot be definitely said that Cl4—1abeled organic acids are released by the kidney (Results, Section III). As is the case with the amino acids, the kidney appears to release very little (if any) organic acids into the renal venous blood. The data presented here are in agreement with other findings that the kidney has a 109 net uptake rather than production of citrate, lactate, pyruvate, and d—ketoglutarate (Cohen, 1964). These latter studies measured only net uptake or production of the organic acids; thus, there still may be movement of organic acids into the renal venous blood although net uptake from blood perfusing the kidney is occurring. For example, Chinard gt g1. (1962) have presented evidence that lactate is formed as well as utilized by the 1t 3132 dog kidney. Kidney tissue, on the other hand, did contain C14- labeled aspartic acid, succinate, malate, and citrate and/or pyruvate. It seems that Cl4—glutamine is con- verted to Cl4-g1utamic acid (present in significant amounts in all 3 kidneys) which then enters the tri- carboxylic acid (TCA) cycle. Although some TCA cycle intermediates are clearly labeled within the kidney, none of these appear in significant quantities in the renal venous blood with the possible exception of fumarate and citrate. When these experiments were proposed, the Cl4 activ— ity in the kidney and the Cl4 activity in the degradation products of Cl4—glutamine could only be surmised. It is apparent from the amino acid and organic acid data that if amino and organic acids other than those described above are labeled in kidney and blood samples each amino 110 14 activ— or organic acid contains less than 0.10% of the C ity injected. The experiments performed and described in the Re- sults show that Cl4-glucose was produced from C14— glutamine by all 3 kidneys and was released into the renal venous blood in increasing amounts over the 5 minutes studied. This was especially notable in Experiment I where C14 activity in the amino and organic acid fraction steadily decreased from blood sample 1 to 5, but C14- glucose activity increased 20-fold during this time. Cl4—glucose was also measured in renal tissue but the total amount was less than that found in all 5 blood samples which indicates that glucose being formed within the intact kidney is rapidly released into the blood. The C14 content of renal glycogen was not assayed in these studies. Cl4—glucose production by the kidney has been found in 2 other independent investigations. Roxe gt g1. (1970) injected tracer amounts of Cl4-glutamine into the renal artery of the dog and found Cl4—glucose appearing in the renal venous blood within one minute after the injection had been made. Cl4—g1ucose was assayed by isolation as phenylglucosazone. Selleck (personal communication) in- jected Cl4—a-ketoglutarate (a degradation product of glutamine) into the renal artery and found evidence for Cl4—glucose in the renal venous blood. Glucose in these 111 latter experiments was separated by water-butanol parti— tion chromatography with a Celite column. Selleck found no significant Cl4 activity in any organic acid (which was not originally present in the injection solution) in the renal venous blood samples through the fifth minute and also found Cl4—aspartic acid, —ma1ate, —succinate, and -a—ketoglutarate within the kidney. The following conclusions are indicated by the studies in this thesis and that of Roxe gt g1. (1970). Cl4—glutamine extracted by the kidney can produce C14— glucose of which more is added to renal venous blood than remains in the kidney. Labeled amino and organic acids, on the other hand, are not significantly added to the renal venous blood within 5 minutes after the injection of Cl4—glutamine into the renal artery. Since acute whole body acidosis can stimulate renal glucose production and since glutamine can be used as a substrate for renal gluconeogenesis, the animal's acid-base state may directly or indirectly regulate renal glutamine metabolism. The site or sites of regulation are still uncertain, but glutamine‘s participation in renal gluconeogenesis sug— gests that factors regulating gluconeogenesis (gluco— corticoids, insulin availability, and adenine and pyri- dine nucleotides) may indirectly control glutamine metab- olism. The enzymes glutaminase I and PEP carboxykinase 112 may also be important to the total regulation of glutamine metabolism. To study more thoroughly the path of C14 from gluta— mine, amino acid and organic acid specific activities must be measured at endogenous levels. A technique has been recently described for studies of this nature (Lyon and Pitts, 1969). Secondly, a stable solution of C14— glutamine must be prepared which can be administered to the animal. The significant presence of pyrrolidone 5- carboxylic acid in the injection solutions (Table 15, 16, and 17) indicates degradation of Cl4-g1utamine in these solutions. In order to study the metabolism of extracted C14- l4—1abe1ed compounds should be re- glutamine, all other C moved so that they will not be extracted by the kidney. Thus, any Cl4-1abeled products of renal metabolism can be said to have come from Cl4—glutamine. Since the quan- tity of Cl4—1abeled amino or organic acids released into the renal venous blood is very small, the injection solu- tions must be virtually free of these compounds. SUMMARY AND CONCLUSIONS Net renal glucose production was studied in the 1g g1tg pump—perfused dog kidney at normal blood pH, during acute whole body metabolic acidosis, and during local renal metabolic acidosis. Acute whole body acidosis caused a statistically significant (P < .01) change from a net renal glucose uptake of 0.92 mg/min during a control period to a net renal glucose production of 0.68 mg/min after one hour of acidosis. However, in a different group of dogs,_when only the renal arterial blood was made acidotic and the systemic blood held at normal pH; the net glucose production of 0.31 mg/min measured after one hour of local renal acidosis was not significantly dif- ferent from the net glucose production of 0.32 mg/min measured during the control period. In neither experiment, whole body nor local acidosis, was there a change in renal glycogen levels. The data from these 2 experiments sug- gest that decreased renal pH per se does not cause in- creased renal gluconeogenesis 13 3139. Since renal glu- cose production was measured after only one hour of local acidosis, the conclusion may not be valid for an acute local acidosis longer than one hour. However, the data obtained during acute whole body acidosis do indicate 113 114 that renal gluconeogenesis can be regulated by factors other than local renal pH. When Cl4—glutamine was added to whole blood and the blood incubated at 37° C for 5 minutes, no Cl4-glutamine entered the blood cells. An amino acid chromatogram of the whole blood after incubation showed that a maximum of 7.0% of the Cl4-glutamine was degraded and that no C14- glucose was produced from Cl4—glutamine during this 5 minute incubation. This data suggested that Cl4-glucose production from Cl4-g1utamine by the 1g g1tg dog kidney l4-glutamine injected ‘ could be studied because (1) all C into the renal arterial blood would remain in the plasma and be available for renal extraction and (2) there was 14 no major degradation of C -glutamine by plasma and no formation of Cl4-g1ucose. After Cl4-glutamine was injected into the renal artery of an 13 g1tg pump-perfused kidney, 5 consecutive one—minute renal venous blood samples were collected and the kidney was taken from the body and frozen after 5 minutes. Each blood sample, the kidney tissue, and the injection solution were assayed for Cl4—labeled amino acids, organic acids, and glucose. In the renal venous bloods no Cl4—amino acid or Cl4-organic acid was added by the kidney in amounts detectable by the techniques employed with the possible exception of fumarate and citrate. However, a significant amount of glucose was 115 found in the renal venous blood from all 3 animals (2 animals at normal pH and one animal after one hour of acute whole body acidosis). On the other hand, kidney tissue did contain Cl4 activity in aspartic acid, succinate, malate, and citrate and/or pyruvate. Cl4-glutamine and Cl4—g1utamic acid were also present in the kidney. Cl4-g1ucose was identi— fied but the total Cl4-g1ucose activity was less in kidney tissue than in renal venous blood. From these data it appears that Cl4-g1utamine enters the kidney and is deamidated to C14—glutamic acid. Part of the Cl4—glutamic acid is converted by the kidney to Cl4-glucose which is added to the renal venous blood. The amount of Cl4—glucose produced by the kidney during the 5 minute experiment was greater than the total amount 14 of any single Cl4-amino acid or C —organic acid present in the renal venous blood. However, it cannot be defi— nitely concluded from these data that absolutely no C14— 1abeled amino or organic acid is released by the kidney within 5 minutes after injection of Cl4-glutamine into the E renal arterial blood. There may have been Cl4-amino and Cl4-organic acids produced by the kidney and present in the renal venous blood with Cl4 activity below the sensi- tivity of the methods used (0.10% of the Cl4 activity in— jected). However, the data do show that Cl4-glutamine extracted during normal acid—base balance and during BIBLIOGRAPHY 117 BIBLIOGRAPHY Addae, S. K. and W. D. Lotspeich. Relations between glu— tamine utilization and production in metabolic acidosis. Am. i. Physiol. 215:269-277, 1968. Alleyne, G. A. 0. Concentrations of metabolic inter— mediates in kidneys of rats with metabolic acid— osis. Nature. 217:847—848, 1968. Alleyne, G. A. O. and G. H. Scullard. Renal metabolic re- sponse to acid base changes. I. Enzymatic control of ammoniagenesis in the rat. 3. Clin. Med. 48: 364—370, 1969. — Balagura, S. and R. F. Pitts. Excretion of ammonia in— jected into renal artery. Am. i. Physiol. 203: 11—14, 1962. Balagura-Baruch, S., L. M. Shurland, and T. C. Welbourne. Effects of a—ketoglutarate on renal ammonia release in the intact dog. Am. i. Physiol. 218:1070-1075, 1970. Benoy, M. P. and K. A. C. Elliot. CLVIII. The metabolism of lactic and pyruvic acids in normal and tumor tissues. V. Synthesis of carbohydrate. Biochem. g. 31:1268—1275, 1937. Bergman, H. and D. R. Drury. The relationship of kidney function to the glucose tuilization of the extra abdominal tissues. Am. i. Physiol. 124:279-284, 1938. Bergmeyer, H. V. Methods of Enzymatic Analysis. Academic Press, New York, 1965. Bersentes, T. J. and D. H. Simmons. Effects of acute acidosis on renal hemodynamics. Am. i. Physiol. 212:633—640, 1967. Bignall, M. C., O. Elebute, and W. D. Lotspeich. Renal protein and ammonia biochemistry in NH4Cl acidosis and after uninephrectomy. Am. 1. Physiol. 215: 289-295, 1968. 118 119 Bondy, P, K., D. F. James, and B. W. Farrar. Studies of the role of the liver in human carbohydrate metab— olism by the venous catheter technic. I. Normal subjects under fasting conditions and following the injection of glucose. J. Clin. Invest. 28:238- 244, 1949. _ Bowman, R. H. Gluconeogenesis in the isolated perfused rat kidney. 3. Biol. Chem. 245:1604-1612, 1970. Chinard, F. P., T. Enns, and M. F. Nolan. Indicator— dilution studies with "diffusible" indicators. Circ. Res. 10:473-490, 1962. Churchill, P. C. and R. L. Malvin. Relation of renal gluconeogenesis to ammonia production in the dog. Am. i. Physiol. 218:241—245, 1970a. Churchill, P. C. and R. L. Malvin. Relation of renal gluconeogenesis to ammonia production in the rat. Am. i. Physiol. 218:353—357, 1970b. Clapp, J. R., E. E. Owen, and R. R. Robinson. Contribu- tion of the proximal tubule to urinary ammonia ex- cretion by the dog. Am. 1. Physiol. 209:269-272, 1965. Clark, B. and J. W. Porteous. Determination of succinic acid by an enzymic method. Biochem. g. 93:21c- 22c, 1964. Cohen, J. J. Specificity of substrate utilization by the dog kidney in vivo. In Renal Metabolism and Epidermology of_§ome Renal—Diseases, Edited by J. Metcoff, Maple Press, York, Pa., 1964. Cohn, C., B. Katz, and M. Kolinsky. Renal gluconeogenesis in the intact dog. Am. i. Physiol. 165:423—428, 1951. Davies, B. M. A. and J. Yudkin. Studies in biochemical adaptation. The origin of urinary ammonia as indi— cated by the effect of chronic acidosis and alkalosis on some renal enzymes in the rat. Biochem. g. 52: 407-412, 1952. DeMeutter, R. C. and W. W. Shreeve. Conversion of DL— lactate—Z—Cl4 or pyruvate—Z-Cl4 to blood glucose in humans: effects of diabetes, insulin, tolbutamide and glucose load. 3. Clin. Invest. 42:525-533, 1966. 120 Dzurik, R., B. Krajci—Lazary, and T. R. Niederland. Glu- cose metabolism in rat kidney: influence of insulin and adrenaline. g. Physiol. 168:782-786, 1963. Exton, J. H. and C. R. Park. Control of gluconeogenesis in liver. I. General features of gluconeogenesis in the perfused livers of rats. J. Biol. Chem. 242:2622—2636, 1967. _ Exton, J. H. and C. R. Park. Control of gluconeogenesis in liver. II. Effects of glucagon, catecholamines, and adenosine 3', 5'—monophosphate on gluconeogenesis in the perfused rat liver. J. Biol. Chem. 243: 4189-4196, 1968. — Flinn, R. B., B. Leboeuf, and G. F. Cahill, Jr. Metabolism on C14—labeled substrates in kidney cortical slices from normal and alloxan—diabetic rats. Am. J. Physiol. 200:508—510, 1961. "' ’ Friedmann, B., E. H. Goodman, Jr., and S. Weinhouse. Ef- fects of insulin and fatty acids on gluconeogenesis in the rat. 3. Biol. Chem. 242:3620-3627, 1967. Glabman, S. R., M. Klose, and G. Giebisch. Micropuncture study of ammonia excretion in the rat. Am. 1. Physiol. 205:127—132, 1963. Goldstein, L. and C. J. Kensler. Factors which affect the activity of glutaminase I in the guinea pig kidney. 1. Biol. Chem. 235:1086—1089, 1960. Goldstein, L. Actinomycin D inhibition of the adaptation of renal glutamine—deaminating enzymes in the rat. Nature. 205:1330-1331, 1965. Goldstein, L. Relation of glutamine to ammonia produc- tion in the rat kidney. Am. 3. Physiol. 210:661— 666, 1966. Goldstein, L. Pathways of glutamine deamination and their control in the rat kidney. Am. i. Physiol. 213: 983—989, 1967. Good, C. A., H. Kramer, and M. Somogyi. The determination of glycogen. 1. Biol. Chem., 100:485-494, 1933. Goodman, A. D., R. E. Fuisz, and G. F. Cahill, Jr. Renal gluconeogenesis in acidosis, alkalosis, and potassium deficiency: Its possible role in regulation of renal ammonia production. 1. Clin. Invest. 45:612—619, 1966. 121 Goorno, W. E., F. C. Rector, Jr., and D. W. Seldin. Rela— tion of renal gluconeogenesis to ammonia production in the dog and rat. Am. 1. Physiol. 213:969—974, 1967. Haddy, F. J., J. Scott, M. Fleishman, and D. Emanuel. Ef— fect of change in renal venous pressure upon renal vascular resistance, urine, and lymph flow rates. Am. i. Physiol. 195:97—110, 1958. Hayes, C. P., Jr., E. E. Owen, and R. R. Robinson. Renal ammonia excretion during acetazolamide or sodium bicarbonate administration. Am. J. Physiol. 210: 744—750, 1966. — _ Hummel, J. P. The fluorometric determination of malic acid. 1. Biol. Chem. 180:1225—1228, 1949. Jacobs, M. H. Some aspects of cell permeability to weak electrolytes. Cold Spring Harbor Symp. Quant. Biol. 8:30—39, 1940. Janicki, R. H. and L. Goldstein. Glutamine synthetase and renal ammonia metabolism. Am. 1. Physiol. 216:1107- 1110, 1969. Jungas, R. L. and E. G. Ball. Studies on the metabolism of adipose tissue. XII. The effects of insulin and epinephrine on free fatty acid and glycerol production in the presence and absence of glucose. Biochem. 2:383—388, 1963. Kamm, D. E., R. E. Fuisz, A. D. Goodman, and G. F. Cahill, Jr. Acid—base alterations and renal gluconeogenesis: Effect of pH, biocarbonate concentration, and PCO . 3. Clin. Invest. 46:1172—1177, 1967. 2 Kamm, D. E. and G. F. Cahill, Jr. Effect of acid—base status on renal and hepatic gluconeogenesis in diabetes and fasting. Am. i. Physiol. 216:1207— 1212, 1969. Kamm, D. E. and R. R. Asher. Relation between glucose and ammonia production in renal cortical slices. Am. 3. Physiol. 218:1161—1165, 1970. Krebs, H. A. Metabolism of amino acids. IV. The synthe- sis of glutamine from glutamic acid and ammonia, and the enzymic hydrolysis of glutamine in animal tis— sues. Biochem. g. 29:1951—1969, 1935. 122 Krebs, H. A., R. Hems, and T. Gascoyne. Renal gluconeo— genesis. IV. Gluconeogenesis from substrate combi— nations. Acta Biol. Med. Germ. 11:607-615, 1963a. Krebs, H. A., D. A. H. Bennett, P. deGasquet, T. Gascoyne, and T. Yoshida. Renal gluconeogenesis. The effect of diet on the gluconeogenic capacity of rat-kidney- cortex slices. Biochem. g. 86:22-27, 1963b. Krebs, H. A., R. N. Speake, and R. Hems. Acceleration of renal gluconeogenesis by ketone bodies and fatty acids. Biochem. 1. 94:712-720, 1965. Krebs, H. A. and P. Lund. Formation of glucose from hexoses, pentoses, polyols, and related substances in kidney cortex. Biochem. g. 98:210-214, 1966. Krebs, H. A., R. Hems, M. J. Weidemann, and R. N. Speake. The fate of isotopic carbon in kidney cortex synthe- sizing glucose from lactate. Biochem. g. 101: 242-249, 1966. Landau, B. R. Gluconeogenesis and pyruvate metabolism in rat kidney, 13 vitro. Endocr. 67:744—751, 1960. Lee, J. B., V. K. Vance, and G. F. Cahill, Jr. Metabolism of C14—1abeled substrates by rabbit kidney cortex and medulla. Am. i. Physiol. 203:27-36, 1962. Lee, Y. P. and T. Takahashi. An improved colorimetric determination of amino acids with the use of ninhydrin. Anal. Biochem. 14:71-77, 1966. Lotspeich, W. D. and R. F. Pitts. The role of amino acids in the renal tubular secretion of ammonia. 1. Biol. Chem. 168:611—622, 1947. Lowenstein, J. M. (Ed.). Citric Acid Cycle. Control and Compartmentation. Marcel Dekker, Inc., New York, 1969. Lyon, M. L. and R. F. Pitts. Species differences in renal glutamine synthesis 1m vivo. Am. i. Physiol. 216: 117—122, 1969. Mahler, R. J. and O. Szabo. Metabolic effects of insulin on rat kidney after inhibiting degradation of the hormone. Endocr. 83:1166-1172, 1968. 123 Mahler, R., W. S. Stafford, M. E. Tarrant, and J. Ashmore. The effect of insulin on lipolysis. Diabetes. 13: 297—302, 1964. Mattenheimer, H., V. E. Pollack, and R. C. Muehrcke. Quantitative enzyme patterns in the nephron of the healthy human kidney. Nephron. 7:144—154, 1970. McCann, W. P. and J. R. Jude. The synthesis of glucose by the kidney. Bull. Johns Hopkins HOSE. 103:77-93, 1958. McCann, W. P., O. D. Gulati, and H. C. Stanton. Renal glucose metabolism during diuresis induced by in— fusions of hypotonic saline. Bull. Johns Hopkins Hosp. 108:36-47, 1961. McCann, W. P. Renal glucose production and uptake in separate sites, and its significance. Am. i. Physiol. 203:572-576, 1962. Meister, A. Metabolism of glutamine. Physiol. Rev. 36: 103—127, 1956. Mirsky, I. A. and R. H. Broh—Kahn. The inactivation of insulin by kidney extracts. I. The distribution and properties of insulin inactivating extracts (Insulinase). Arch. Biochem. 20:1-9, 1949. Morris, M. E. and R. A. Millar. Blood pH/plasma cate— cholamine relationships: non—respiratory acidosis. Brit. 3. Anaesth. 34:682—688, 1962. Mortimore, G. E. Effect of insulin on release of glucose and urea by isolated rat liver. Am. i. Physiol. 204:699-704, 1963. Nadkarni, G. B. and K. E. Chitnis. Effect of insulin on gluconeogenesis from glycine—2—Cl4. Arch. Biochem. Biophys. 101:466-470, 1963. Nishiitsutsuji-Uwo, J. M., B. D. Ross, and H. A. Krebs. Metabolic activities of the isolated rat liver. Biochem. g. 103:852-862, 1967. O'Donovan, D. J. and W. D. Lotspeich. Activation of kidney mitochondrial glutaminase by inorganic phosphate and organic acids. Nature. 212:930-932, 1966. 124 O'Donovan, D. J. and C. W. Warner. A quantitative evalua- tion of the non—enzymatic conversion of glutamine to ammonia. Experientia. 25:100-101, 1969. Oelert, H. and W. Nagel. Die Abhangigheit der Ammonia- Produktion von der GFR bei Ureterabklemmung und bie Durchblutungsdrosselung in der Hundeniere. Arch ggg. Physiol. 292:129—139, 1966. Owen, E. E. and R. R. Robinson. Amino acid extraction and ammonia metabolism by the human kidney during the prolonged administration of ammonium chloride. J. Clin. Invest. 42:263—276, 1963. _ Owen, O. E., P. Felig, A. P. Morgan, J. Wahren, and G. F. Cahill, Jr. Liver and kidney metabolism during prolonged starvation. J. Clin. Invest. 48:547— 583, 1969. _ Patrick, S. J. Renal gluconeogenesis, adenine nucleotides, and enzyme activities. Can. 3. Biochem. 46:1345— 1349, 1968. Piez, K. A. and L. Morris. A modified procedure for the automatic analysis of amino acids. Anal. Biochem. 1:187—201, 1960. Pilkington, L. A., T. K. Young, and R. F. Pitts. Proper— ties of renal luminal and antiluminal transport of plasma glutamine. Nephron. 7:51-60, 1970. Pitts, R. F., J. deHaas, and J. Klein. Relation of renal amino and amide nitrogen extraction to ammonia pro— duction. Am. 1. Physiol. 204:187-191, 1963. Pitts, R. F. Renal production and excretion of ammonia. Am. i. Physiol. 36:720-742, 1964. Pitts, Robert. Physiology of the Kidney and Body Fluids. Year Book Medical Publishers, Chicago, 1968. Pollack, V. E. and H. Mattenheimer. Glutaminase activity in the kidney in gout. 1. Lab. Clin. Med. 66:564- 570, 1965. Pollack, V. E. and H. Mattenheimer, H. De Bruin, and K. J. Weinman. Experimental metabolic acidosis: The enzymatic basis of ammonia production by the dog kidney. 1. Clin. Invest. 44:196-181, 1965. 125 Preuss, H. G. Pyridine nucleotides in renal ammonia metab- olism. 3. Lab. Clin. Med. 72:370-383, 1968. Preuss, H. G. Renal glutamate metabolism in acute metabolic acidosis. Nephron. 6:235—246, 1969. Rector, F. D., Jr., D. W. Seldin, and J. H. Copenhaver. The mechanism of ammonia excretion during ammonium chloride acidosis. 3. Clin. Invest. 34:20—26, 1955. Rector, F. D. and J. Orloff. The effect of the administra- tion of sodium biocarbonate and ammonium chloride on the excretion and production of ammonia. The absence of alterations in the activity of renal ammonia- producing enzymes in the dog. 1. Clin. Invest. 38: 366-372, 1959. Reinecke, R. M. The metabolism of fructose by the eviscer- ated rat. Am. 1. Physiol. 136:167—172, 1942. Reinecke, R. M. The kidney as a source of glucose in the eviscerated rat. Am. i. Physiol. 140:276-285, 1943. Reinecke, R. M. and S. Roberts. The effect of fasting on the blood sugar curve of the eviscerated rat. Am. 3. Physiol. 141:476-479, 1944. Reinecke, R. M., G. G. Rudolph, and M. J. Bryson. Effect of ureteral ligation, phloridizin and mercury bi— chloride on the glucogenic function of the kidney. Am. 3. Physiol. 151:198-201, 1947. Reinecke, R. M. and P. J. Hauser. Renal gluconeogenesis in the eviscerated dog. Am. i. Physiol. 153:205— 209, 1948. Roberts, S. and L. T. Samuels. Fasting and gluconeo— genesis in the kidney of the eviscerated rat. Am. 3. Physiol. 142:240—245, 1944. Roxe, D. M., J. DiSalvo, and S. Balagura-Baruch. Renal glucose production in the intact dog. Am. 1. Physiol. 218:1676-1681, 1970. Russell, J. A. and A. E. Wilhelmi. Glyconeogenesis in kidney tissue of the adrenalectomized rats. A. Biol. Chem. 140:747—754, 1941. 126 Sartorius, O. W., J. C. Roemmelt, and R. F. Pitts. The renal regulation of acid—base balance in man. IV. The nature of the renal compensations in ammonium chloride acidosis. 1. Clin. Invest. 28:423-439, 1949. Sayre, F. W. and E. Roberts. Preparation and some prop— erties of a phosphate-activated glutaminase from kidneys. 1. Biol. Chem. 233:1128~1134, 1958. Schimmel, R. J. and E. Knobil. Role of free fatty acids in stimulation of gluconeogenesis during fasting. Am. 1. Physiol. 217:1803—1808, 1969. Schimmel, R. J. and E. Knobil. Insulin, free fatty acids, and stimulation of hepatic gluconeogenesis during fasting. Am. 1. Physiol. 218:1540-1547, 1970. Selleck, B., J. J. Cohen, and H. M. Randall, Jr. Enzymic assay for a-ketoglutarate in dog blood, plasma, urine, and tissue. Anal. Biochem. 7:178-188, 1964. Shalhoub, R., W. Webber, S. Glabman, M. Canessa—Fischer, J. Klein, J. deHaas, and R. F. Pitts. Extraction of amino acids and their addition to renal blood plasma. Am. 1. Physiol. 204:181—186, 1963. Simpson, D. P. and D. J. Sherrard. Regulation of glutamine metabolism 13 vitro by bicarbonate ion and pH. 1. Clin. Invest. 48:1088-1096, 1969. Séling, H. D., B. Willms, D. Friedrichs, and J. Kleineke. Regulation of gluconeogenesis by fatty acid oxida- tion in isolated perfused livers of non-starved rats. 131. 1. Biochem. 4:364-372, 1968. Steiner, A. L., A. D. Goodman, and D. H. Treble. Effect of metabolic acidosis on renal gluconeogenesis 1m vivo. Am. 1. Physiol. 215:211-217, 1968. Stone, W. J. and R. F. Pitts. Pathways of ammonia metab- olism in the intact functioning kidney of the dog. 1. Clin. Invest. 46:1141—1150, 1967. Sullivan, L. P. and M. McVaugh. Effect of rapid and transitory changes in blood and urine pH on NH4 excretion. Am. 1. Physiol. 204:1077-1085, 1963. Teng, C. T. Studies on carbohydrate metabolism in rat kidney slices. 1. Metabolism of glycerol and pyru— vate. Arch. Biochem. Biophys. 48:409-414, 1954a. 127 Teng, C. T. Studies on carbohydrate metabolism in rat kidney slices. II. Effect of alloxan diabetes and insulin administration on glucose uptake and glucose formation. Arch. Biochem. Biophys. 48:415—423, 1954b. Usatenko, M. S. Hormonal regulation of phosphoenol— pyruvate carboxykinase activity in liver and kidney or adult animals and formation of this enzyme in developing rabbit liver. Biochem. Mgg. 3:298-310, 1970. VanSlyke, D. D., R. A. Phillips, P. B. Hamilton, R. M. Archibald, P. M. Futcher, and A. Hillar. Glutamine as source material of urinary ammonia. 1. Biol. Chem. 150:481-482, 1943. Weber, G., R. L. Singhal, and S. K. Srivastava. Insulin: Suppressor of biosynthesis of hepatic gluconeogenic enzymes. Proc. Nat. Acad. Sci., U.S. 53:96-104, 1965. Weidemann, M. J., D. A. Hems, and H. A. Krebs. Effects of nucleotides on renal metabolism. Nephron. 6: 282—296, 1969. Weiss, R. F. and H. G. Preuss. Glutamine synthetase and plasma glutamine in augmented ammoniagenesis in acidosis. Am. 1. Physiol. 218:1697—1700, 1970. White, L. W. and B. R. Landau. Effect of glucocorticoids on metabolism of carbohydrate by kidney cortex. Am. 1. Physiol. 211:449—456, 1966. Williamson, J. R., E. T. Browning, and R. Scholz. Control mechanisms of gluconeogenesis and ketogenesis. I. Effects of oleate on gluconeogenesis in perfused rat liver. 1. Biol. Chem. 244:4607—4616, 1969. Yoshida, T., C. Cohn, and Y. H. Maa. Effects of adrenal- ectomy and manner of food intake on renal gluconeo- genesis. Endocr. 84:417-420, 1969. APPENDIX 128 FREQUENTLY USED ABBREVIAT IONS PFF protein-free filtrate PAH p—aminohippuric acid TCA cycle tricarboxylic acid cycle Tm maximum tubular transport rate PSI pounds per square inch SE standard error I.S. injection solution a—KG a—ketoglutarate [ ] concentration per liter of solution mOs milliosmole mg % mg/100 ml of solution mM millimolar uM micromolar Kg kilogram 9 gram mg milligram pg microgram wt weight ml milliliter L liter mu millimicron (wavelength) mc millicurie 129 hhhh ulflflfllfljfifisufirflflinjfl171111111111“