Mutarotase and glucose anomers

lUOCHEMIC.\L

1, 6246

MEDICINE

(1067)

Mutarotase

and

Glucose

Anomers

1. The Contribution of Various Organs to the in Vivo Mutarotation of Exogenous Glucose and the Clearance of Glucose Anomers by the Kidney of the Anesthetized Dog

JOHN II. Mutarotase

JOHN

B. HILL,

B. HILL Activity

DORIS

AND

S. COWART

in Chemically Rat Kidneys

T. U. L. BIBER,l

AND

Damaged

WILLIAM

Glycosuric

D. HUFFINES

Departments of Pharmacology, Medicine and Pathology, University of North Carolina School of Medicine, Chapel Hill, North Carolina .%7614 Received

May

4, 1967

Although glucose is known to exist in at least two anomeric forms (Fig. 1) until recently (1) nothing was known of the state in which glucose exists in biological fluids. The concept that the various forms of glucose might play some role in health and disease is not a new one. Lundsgaard and Holboll (2) in the 1920s published a series of papers in which a “new-glucose” present in the blood of nondiabetic patients was found to be lacking in diabetics. It could be restored to the blood of diabetics by the administration of insulin and could be produced in vitro by combining insulin, muscle tissue, and ordinary glucose. Its presence was measured by a change in optical rotation of diffusates of blood or glucose-insulin-muscle mixtures in the absence of a change in reducing power. These papers make fascinating reading, but the work could not be confirmed by a substantial number of the author’s contemporaries. In 1949 Bentley and Neuberger (3) working with glucose oxidase from a mold, found evidence that certain preparations contained a substance which catalyzed the mutarotation of glucose. The mutarotational catalyst was separated from the glucose oxidase in 1952 by Keilin and Hartree ’ Present sachusetts.

address

Biophysical

Laboratory,

Harvard 62

Medical

School,

Boston,

Mas-

MUTAROTASE

AXD

GLUCOSE

63

ANOMERS

(4) who established its enzymic properties and named it mutarotase. In 1954 Keston (5) reported finding mutarotase in mammalian tissues and proposed a theory of sugar transport involving that enzyme. He further suggested an involvement of mutarotase in diabetes mellitus and renal diabetes. Bailey, Fishman, and Pentchev (6) reported mutarotase to be present in certain plants. Wallenfels and Herrmann (7) found mutarotase in E. coli. MUTAROTAAION CH,OH

CH,OH

H(/qpEKT>H(ry

OH

OH

a-D-Glucose

D-D-Glucose

36%

64% FIQ.

1. The

mutarotation

of

glucose.

Enzymes which catalyze the mutarotation of glucose and other monosaccharides have now been reported to be present in plants, molds, bacteria, fish, amphibia, birds, and mammals (4-8). The purpose served by such enzymes has not been elucidated. Other enzymes specific for a particular anomer of glucose or glucose-6-phosphate, as well as enzymic reactions one product of which is a particular anomer of glucose have been described (4, 9, 10, 11). This suggests that there may be instances when spontaneous mutarotation is rate limiting. The presence, then, of an aldose-1-epimerase (mutarotase) could circumvent the rate limitation. Keston (8) has collected evidence which he feels is consistent with “the control of the limiting rate of carbohydrate metabolism and transport by mutarotase”. In an attempt to elucidate a possible relationship to disease states in man, we have begun to study tissues obtained at autopsy for mutarotase activity. Although that study is not yet ready for publication, preliminary findings have clearly shown that correlations between tissue levels and certain disease states can be made. For example, pyelonephritic kidneys have low mutarotase activity in the renal cortex. Two other less firmly established findings are low muscle mutarotase activities in a high proportion of the diabetics in our series, and the highest liver enzyme activity recorded was from a four and one half month old infant with glycogenosis Type II. The experiments which follow are attempts to learn more of the role

64

HILL,

COWART,

AND

HILL,

BIBER,

.iSD

HUFFINES

played by the glucose anomers and mutarotase (aldose-1-epimerase) in the biochemistry of the mammalian organism. The finding of catalysis of a particular reaction in vitro is no guarantee that the same reaction will be catalyzed in vivo. Data previously published (1) show that cY-n-glucose infused into the blood stream of the intact anesthetized dog is brought to equilibrium more rapidly than a-n-glucose infused into a Starling heart-lung preparation, or added to shed dog blood. The experiments to be described in the first part of Part I of this communication extend these observations and indicate that the liver, an organ rich in mutarotase, accelerates the attainment of equilibrium more than other organs studied. The highest concentrations of mutarotase activity have been found in the kidney and phlorizin, which interferes with glucose reabsorption by the renal tubule, inhibits mutarotase (5). The aglomerular fish which does not reabsorb glucose from kidney tubule has very little mutarotase activity in its kidney (Sj. These findings prompted the study of glucose anomer clearance by the kidney which comprise the second portion of Part I of this communication. Potassium dichromate, mercuric chloride, and uranyl nitrate have long been known to produce glycosuria when injected into laboratory animals in sufficient amounts (12). Hepler and Simonds (13) used t.hese salts to produce glycosuria in dogs to study the kidney cortex alkaline phosphatase activity in an attempt to shed some light on a possible role of phosphatase in the reabsorption of glucose by the kidneys. Kidney alkaline phosphatase activity was reported to be decreased by potassium dichromate, by mercury bichloride, but not by uranyl nitrate. Keston’s evidence (8) suggesting that mutarotase may play a role in glucose transport, and his finding the enzyme to be plentiful in the kidneys of all species studied (with the exception of the aglomerular fish) prompted this study of the mutarotase activity of the kidneys of animals treated with glycosuria producing quantities of these same salts. These experiments are described in Part II of this communication. PART

I MATERIALS

AND

METHODS

The method for determining deviations from anomer equilibrium (36% Q and 64% P-n-glucose) used in this study utilizes a continuous sampling technique and has been fully described in a previous publication (1). The principle of the method, schematically represented in Fig. 2, is based upon the specificity of glucose-oxidase for the p anomer of n-glucose and the catalytic action of hydroxyl ions on spontaneous mutarotation. The blood sample to be analyzed is dialyzed and t,he glucose which passes

through the dialyzer membrane is divided into two portions. Each portion is analyzed for /3-n-glucose, one with and one without equilibration of the anomers with alkali. The analysis is standardized with equilibrium glucose solutions. With this procedure any deviation from anomer equilibrium appears as a difference (A) between the alkali equilibrated portion, the NaOH value, and the nonequilibrated portion, the NaCl value. These differences are plotted at the bottom of Figs. 3, 4, and 5 along the abscissa. Experiments were done in mongrel dogs of either sex, fed for the last time at 4 pm the day prior to the experiment, anesthetized with phenobarbital, and heparinized as previously described (1). SCHEMATIC REPRESENTATION OFa,f3 GLUCOSE ANALYSIS SRl4tPL E

Waste

t-

i%ziE A L A’

RECfPiENT NaOH NaCL ,&WCOSE REAGENTS BUFFERED

--,

AL

(

=

\

-

AA A ‘-A

e L. B’

NaCL Value Coiorimefer NaOH Value Cdorhefer

,L ,, Two Pen Recorder

Refrigerated Bath5“ t?=Dia~zed t NaCL -wo change B’= Diafyzed t NaOH -wquilirmm

FIG. 2. Schematic representation

of technique

for detecting deviations from glucose

anomer equilibrium. Glucose infusions were given at a rate of 2 ml/minute by means of a 0.073 inch inside diameter pumping tube on the AutoAnalyzer pump platen. All infusions were made to deliver 20 mg/kg minute of glucose and were dissolved to the appropriate concentration in cold water and kept in ice to minimize spontaneous mutarotation. Infusions were continued long enough so that the deviation from equilibrium, A, in the sampled stream could reach a steady state. This usually occurred within five minutes. The infusion deviation, Ai, was compared with the appropriate pre- and/or postinfusion deviation, A,, and the mean change in deviation, Ai-Ao, estimated. Continuous sampling and infusions were accomplished through the use of polyethylene tubing (PE-60 Clay-Adams, Inc., New York). In the experiments in which samples were drawn from branches of the hepatic vein, the sampling tube was given the requisite rigidity for guidance by inserting it 9.5 cm into a 10 cm sleeve of Teflon tubing (0.047

66

HILL,

COWART,

AXD

HILL,

BIBER,

inch i.d., Bel-Art Products, Pequannock, in the portion of the Teflon tube below with a hot needle. These were for the when contact was made with the vessel liver.

AND

HUFFINES

N. J.) . Several holes were made the tip of the polyethylene tube purpose of preventing occlusion wall within the substance of the

280-7 270260250240230220210200190-

l:bO

I:30

2:00

330

4:oo

4:30

FIG. 3. The mutarotation of an excess of a-n-glucose by the kidney and hind limb of the anesthetized dog. Glucose was infused into the jugular vein.

To determine the role of various parts of the body on the mutarotation of exogenously infused a-D-ghlcose the mean change in deviation caused by the infusion, AI-Ao, was determined in the affluent and effluent blood supply of the region studied. Fig. 6 is a diagramatic representation of the vascular routes to and

MUTAROTASE

AND

GLUCOSE

67

ANOMERS

from the regions studied. All glucose infusions in the first experiments were made into the jugular or femoral vein. Blood was sampled from the femoral artery or the abdominal aorta by means of a tube inserted through a branch of the femoral artery. This blood was considered to be representative of the affluent supply to all regions studied. The effluent blood from the various regions was obtained as follows: A. Head: sampling was accomplished from a tube placed in the jugular vein opposite to the one being used for the infusion; B. Hind limb: sam2302202lO200190B

ISO-

2 9 o s

170-

2

140-

8 7

130-

160150-

$ 2 25

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+J

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IIOIOO60706050f HEPLLTIC VEIN SAMPLING

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FEMORAL ARTERY SAMPLING

l

HEPATIC VEIN SAMPLING

P 2 8 IO ‘\

20

7. I30

:.

1, 200

230

300

I 330

I 400

*-,I 4 30

500

*

P p

FIG. 4. The mutarotation of an excess of cy-n-glucose by the liver. Glucose was infused into the femoral vein. The second curve shows the rise in glucose in the hepatic vein subsequent to an intravenous injection of glucagon.

pling was accomplished from a tube placed in the femoral vein by way of a small venous branch; C. Spleen, stomach and intestines: sampling was accomplished from a tube placed in the hepatic portal vein by way of a branch of the splenic vein; D. Kidney: sampling was accomplished from a tube placed in the left renal vein by way of the left testicular vein; and E. Liver, spleen, stomach, and intestines: sampling was accomplished from a tube placed in a branch of the hepatic vein. For this procedure the dog’s chest and abdomen were opened with a midline inci-

68

HILL,

COWART,

AND

HILL,

HIRER,

AND

HCFFISES

sion. Respiration was supported by means of a tracheal cannula and a positive pressure respirator. The Teflon sleeved sampling tube, previously described, was inserted into the superior vena cava by way of the right jugular vein. It was fed into the inferior vena cava just above the diaphragm where it could be palpated in the lumen. By manipulation of the tube through the wall of the inferior vena cava, the tip could be inserted into a branch of the hepatic vein and from the abdominal cavity could, in some experiments, be felt in the substance of a lobe of the liver. Once 200

-

190180170160I50 -Q 9Q 140Q z 0 c f

130-

8-8

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;

IZOIIO-

so$

80-

:

70-

2

60-

2

50-

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Hcpatic portal infusion

40-

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2Omg/kqmin

2Om9 /k9 min a- D-9lucose I I

3 $ 0

Femoral vein infusion

”

30-

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20IO-

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12730

I:00

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200

30

2 D

20

s

@ v

rh?

FIG. 5. The mutarotation of a-D-ghCOEe infused into the liver by way of the hepatic portal vein. Samples were taken continuously from the femoral artery. The second infusion was the same cY-n-glucose solution infused into the femoral vein.

located, the tube was held in place by means of a clamp placed on the inferior vena cava in such a way as to hold the Teflon tube and yet not occlude the blood flow from the abdominal vena cava and hepatic veins into the heart. About thirty minutes were allowed for clotting before heparinization of the dog. The position of all sampling tubes was verified at post mortem. In a second series of experiments, assessing the role of the liver in

MUTAROTASE

Ah-D

GLUCOSE

Ah-OMERS

69

mutarotation, infusions of glucose were made directly into the hepatic from the abdominal aorta. portal yein. Sampling was accomplished Control infusions were placed in the femoral vein. Estimates of the mutarotase activity in the tissues were made by a previously described method (14). The total mutarotase activity in organs with a heterogeneous distribution of enzyme were obtained by grinding the entire organ in a meat grinder and mixing the resultant

TESTICULAR VEIN

BIG.

6. Vascular routes showing sites of sampling and infusion.

grindings until homogeneity was accomplished as evidenced by a homogeneous enzyme activity content. The total wet. weight of the grindings in gm X units/gm of the sampled portion gives the total unitage. This was done with the stomach and small and large intestines after removal of their contents by opening the lumen and washing out gently with tap water. The entire kidney, heart, lungs, and brain were homogenized in the same manner. Skeletal muscle, liver, and spleen are homogeneous in their enzyme activity content and were therefore not ground in their entirety. Representative samples were taken for assay and the wet weight

70

HILL,

COWART,

AND

HILL,

BIBER,

.~SD

IIITFFISES

of the entire organ was obtained, or in the case of skeletal muscle estimated. Plasma glucose and urine glucose values for the renal clearance studies were determined by automated methods which were modifications of the anomer procedure. Potassium oxalate was used as the anticoagulant. Plasma and urine creatinine were determined by the method of Chasson et al. (15). Urines were diluted at least twenty-fivefold with saturated benzoic acid immediately after collection. Plasmas were diluted with saturated benzoic acid or distilled water to an appropriate concentration just prior to analysis. Bloods were centrifuged immediately after collection and the plasma kept frozen until analyzed. Diluted urines were refrigerated until analyzed. Catheters were placed in both ureters to facilitate urine collections. Creatinine infusions (1.5 mg/ml) were made into the patent jugular vein at a constant rate of from 2-4 ml/minute by means of a pump tube on the Auto-Analyzer pump. Phlorizin 0.2 mg/ml in saline was administered at constant rates in the same manner. Glucose anomers were infused into the aorta rostra1 to the level of the renal arteries by means of a catheter fed through the femoral artery. Continuous blood sampling was accomplished from a catheter placed in the aorta so that it sampled blood caudal to the renal arteries. Intermittent samples were obtained from the femoral artery during the middle of the urine collection period. The clearance experiments were conducted so that periods in which an excess of a-n-glucose, over that seen at thermodynamic equilibrium, might be compared with periods in which an excess of &n-glucose was entering the kidney. Equilibrium glucose infusion periods were also used as noted. The periods were compared in respect to the glucose reabsorbed from the renal tubule. The continuous monitoring of the blood for deviations from anomer equilibrium was used only as a means of being certain that one or the other anomer was entering the kidney in an excessive amount. It was not used for purposes of calculation. The assumption was made that the anomer excess measured caudal to the level of the renal arteries was at least qualitatively representative of the blood entering the renal arteries. This assumption is supported by the finding in another series of experiments (not as yet completed) that the anomer in csccss in the infusion is also in excess in the collected urine. Phlorizin was administered as noted to decrease the reabsorption of glucose and to determine its effects on the results. RESULTS In Fig. 3 are the results of an experiment from which the mean changes in deviation from anomer equilibrium in arterial and venous blood per-

MUTAROTASE

AND

GLUCOSE

ASOMERS

71

fusing the kidney and the hind limb were estimated. As may be seen, the in the femoral artery is approximately haived in the renal and the femoral veins. The slightly larger A seen in the femoral artery at 1:30 pm, as compared to that seen at 4:00 pm, is attributed to spontaneous mutarotation in the iced a-D-ghlCOSe infusion reservior. The break in the curves occurs when the site of the continuous sampling is transferred. Table 1 contains a summary of data obtained from experiments performed on ten dogs which evaluate five separate regions of the body. Though there appears to be a relationship between the sex of the dog and the arterial &-&, this was not found when a larger group of dogs used for other experiments was examined. There is no relationship between the weight of the dog and the arterial 4*-A0. From the number of experiments reported in Table 1, no statistically significant differences in the ability to mutarotate exogenous cu-D-glucose may be seen between regions A-D, even though variation in total mutarotase activity concentration is as great as an order of magnitude. While it is possible that with a greater number of experiments some statistically significant differences might emerge, the within-region variation coupled with the small between-region differences suggested that the information to be gained did not warrant further efforts in this direction. Region E, containing the liver, is however, clearly significantly different from regions A-D. No excess of a-D-glucose was detected in blood drawn from the hepatic veins. The results of one of the hepatic vein sampling experiments may be seen in Fig. 4. 90 significant A appeared during the a-n-glucose infusion until the site of sampling was transferred to the femoral artery. This particular dog (132) differed from the other studied in that the concentration of glucose in the hepatic vein was close to, or lower than, that in the femoral artery. Whether the liver was adding glucose to the circulation (dog 131) or not (dog 132), exogenously administered cu-D-glucose was brought to equilibrium before appearing in the hepatic veins. Glucagon was administered and endogenous glucose was released into the hepatic veins. This glucose is at, or near, equilibrium as was previously demonstrated on sampling from the vena cava in the vicinity of the hepatic veins (1). There is in this experiment, as well as in those previously described, a suggestion that an excess of a-n-glucose might, be released. Validation of this point must await further experimentation because of the modest changes in A which have occurred. To assess further the ability of the liver to mutarotate glucose, infusions of nonequilibrium 4 obtained

glucose were made into the hepatic portal vein while continuously Sampling from the femoral artery. Control infusions were made into the femo-

Hind

B.

limb

E.

(left)

0 Femoral b Estimated

stomach,

and

and

OS

TES

12.8 10.6 11.7 11.3

11.2 12.8 15.6 18.6 11.3 13.6 10.6 11.2

M 26.0 M 20.5

M M M M

125 119 126 127

131 132

F M F M M F M F

122 125 133 118 127 128 119 122

Dog number, sex and weight ill kg.

PERFORMED

artery or abdominal aorta. the weight of muscle in region.

Liver, spleen, intestines

1). Kidney

(1. Spleen, stomach, intestines

Head

;\.

Region

~IWRIXIESTS

23 14

20 15 18 22

13 17 12 17 18 13 14 12

Arteriale Ai-Ao

1

~VALU.lTE

TABLE \17~~~~*

0 0

14 6 13 11

12 12 10 13 9 11 s 9

Venous Ai-A\0

Mean change in A during 20 mg/kg minute infusion (ma/l00 ml)

DOGS

I-

SEPARATE

kterial

Venous

100 100

30 60 2s 50

8 ‘29 li 24 50 15 43 ‘25

(%,

At-A,,

A,-A0

% Mutarotated

FIVE

>

x

100

I:EGIOSS

THE

BODY

wet

0.01

0.20

and intestines

Liver 0.30 Spleen O.o:! Stomach 0.07

Kidney

and intestines

Spleen 0.02 Stomach 0.07

Muscle

Brain 0.10 Muscle 0.01

(units/gm

weight

Mutarotase Activit) Concentratjion

OF

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MUTAROTASE

AND

GLUCOSE

73

ANOMERS

ral vein. The results of a typical experiment may be seen in Fig. 5. No excess of a-n-glucose reached the sampling tube when infused into the hepatic portal vein. When the a-n-glucose was infused into the femoral vein the expected A appeared. The rise and fall curves of the two infusions are nearly superimposable. The slight difference in peak height of the two curves is probably owing to hepatic glucose uptake. Within the error of the method (C 30 seconds) no delay in glucose rise was noted when glucose was infused into the hepatic portal vein as compared with the femoral vein. That this phenomena is reproducible and applies as well to ,&n-glucose may be seen in Table 2. In three of the dogs small excesses of a-n-glucose were seen to reach the arterial circulation.

COMP.~ISOXS

Dog

Kumber 111 112 113 116 lli

OF THE MUTAROTATION VEIN WITH GLUCOSE Glucose hnomer Infused 20 mg/kg minute a CY 8” C-Y Ly

TABLE ‘1 OF GLUCOSE INFUSED INTO THE HEPATIC ISFCSED INTO THE FEMOR.\L VEIN Mean

Change

Portal

Vein 3 4 0 0 0 2

From Infusion

Baseline Femoral

PORTAL

A (mg/lOO

ml)

Vein Infusioll IS ‘)-.,‘, 11 --s 19 10

Fig. 7 shows the results of continuously monitoring for anomer deviations during a renal clearance experiment. It is apparent that the infusion of a single anomer into the dog’s blood stream results in samples containing an excess of that anomer over the amount found at thermodynamic equilibrium. This type of tracing is evidence that an excess of one anomer was indeed entering the renal circulation. Table 3 shows the results in tabular form from an experiment in a nonphlorizin treated dog. No consistent effect of the type of anomer being infused on the percent of the filtered load being reabsorbed is evident. Period 4 and 5, in which different anomers were infused are more alike than any two periods in which the same anomer was infused. Fig. 8 shows a graphical representation of a similar experiment. Here all the points, whether representing periods of cyor j3-n-glucose infusion seem to fall on the same curve. Table 4 shows the results from an experiment in which phlorizin was infused at a rate which decreased the tubular reabsorption of glucose as evidenced by a decrease in reabsorption over the data seen in Table 3 and a decrease over the phlorizin free period. If one compares periods 6-9

74

HILL,

Q

320-

8

310-

s

300-

COWART,

AND

HILL,

BIBER,

Sh-D

HUFFINES

, -ANOMERS ---ANOMERS

EQUILISRATEO WITH ALKAL, NOT EOUILIBRATEO

FIG. 7. Plot of data showing anomer deviation from thermodynamic during infusion of (Yand /3-n-glucose in a renal clearance experiment.

equilibrium

when the plasma glucose levels are reasonably constant, the anomer infused does not appear to influence reabsorption. A similar experiment is seen in Fig. 9 in which periods of CT,p, and equilibrium n-glucose are all plotted on the same graph. Again all the points fall on the same curve. TABLE DOG

104

14.1

Glomerular Collection Filtration period Rate 20 minutes ml/period 1 2 3 4 5 6

1100 1014 951 1210 1000 869

KG

FEMALE-GLUCOSE

Plasma Filtered load glucose mg/ml mg/period 3.10 4.20 4.00 4.70 .5.00 5.30

3410 4259 3804 5687 5000 4518

3 IKFUSEU

350

Glucose Glucose excreted reabsorbed mg/period mg/period 1625 1765 1758 2400 2130 2015

1785 2494 2046 3287 2870 2503

MC:/JIINUTE

‘% Filtered load Anomer reabsorbed infused 52 3 58.6 53.8 57,s 57.4 55.4

a a a! P a 13

MUTAROTASE

AIiD

GLUCOSE

75

ANOMERS

DOG 103 8.9Kg MALE 0 a-D- glucose infusion @p-Dglucose infusion l 0

0

I

I

90

100

I

I

I

I

1

110

120

130

140

150

GLUCOSE FILTERED LOAD mg /min FIG. 8. Glucose reabsorbed versus glucose filtered load in a normal anesthetized dog. Dog 103-3.9 Kg male. 0 a-n-glucose infusion; l P-n-glucose infusion.

DISCUSSION

The passage of blood glucose through the chambers of the heart and the lungs contributes little to mutarotation. It takes approximately the same length of time to attain anomer equilibration in the Starling heartlung preparation as it does in shed dog blood at 37” gassed with 95% O2 TABLE D0~10012.7

Glomerular Collection Filtration period Rate 20 minutes ml/period ~ 1

1135

Plasma Filtered glucose load mg/ml mg/period 2.60

Phlorizin 1165 1172 1052 1041 1090 989 1048 1048

4

KGMALE-GLUCOSEINFUSED

2.70 2.65 2.80 3.00 3.00 3.10 3.10 3.05

2951

200 MG/MINUTE

Glucose excreted mg/period 907

Glucose $%cFiltered reabsorbed Anomer load mg/period reabsorbed infused 2044

69.2

Q

52.6 32.3 29.3 34.4 28.4 26.9 27.2 28.0

a a a

infusion started 0.016 mg/minute 3146 3106 2946 3123 3270 3066 3249 3196

1490 2101 2083 2050 2340 2240 2365 2300

1656 1005 863 1073 930 826 884 896

a* 8* a

76

HILL,

5

AKD

HILL,

BIBER,

DOG 98 PHLORIZIN TREATED 0 a -D-glucose infusion l P-D-glucose infusion c) Equilibrium glucose infusion

25

\F 3 20

COWAR’I,

13.5Kg

AXD

HCFFISES

FEMALE a

i

0 40

, 45

, 50 GLUCOSE

, , 55 60 FILTERED

, , 65 70 LOAD mg/mii

, 75

, 80

FIG. 9. Glucose reabsorbed versus glucose filtered load in a phlorizin treated anesthetized dog. Dog 98 phlorizin treated-13.5 Kg female. 0 a-n-glucose infusion; l P-n-glucose infusion ; and @ equilibrium glucose infusion.

and 5% CO, shaken on a Dubnoff shaker (1). In addition, it was found using the heart-lung preparation that as much mutarotation took place in the glass and plastic tubing of the external circuit as in the dogs heart and lungs (unpublished data). The total mutarotase content of the combined heart and lungs is approximately ten units. The data presented shows that the liver is the most effective organ for restoring glucose anomer equilibrium in blood. This organ is also the richest in mutarotase containing approximately 110 units. The A, or excess of one anomer over that found in the equilibrium glucose mixture obtained when a solution, primarily of one anomer, is infused into the intact dog, reaches a maximum usually within five minutes after the infusion is started. It maintains this maximum level as long as the infusion continues and disappears within five minutes after the infusion is terminated. This phenomenon has been seen repeatedly in the past (1) and may be seen in the present experiments in Figs. 3, 4, and 5. By what mechanisms does this come about? Since the phenomenon occurs whether Q or p-n-glucose is infused (l), selective removal of the anomer in excess of equilibrium quantities seems highly unlikely. Spontaneous mutarotation (1) and glucose turnover (16) have been shown to be too slow to contribute substantially to the rapid reestablishment of anomer equilibrium seen in the intact dog. Dilution of the added glucose in the glucose space must play some role

MUTAROTASE

AND

GLUCOSE

AKOMERS

77

in decreasing the A. The very fact that the A reaches a steady state while the glucose levels are still rising and has disappeared long before the glucose levels return to preinfusion values, dissociates anomer equilibration and glucose deposition with respect to time. This argues against the A being only a function of dilution in the glucose pool. The above cited factors have led to the hypothesis that the reestablishment of anomer equilibrium in the intact dog is probably enzymically catalyzed. It this is true, then the glucose passing through organs which contain mutarotase must in some manner contact the enzyme. In liver, an organ rich in mutarotase and reported to be freely permeable to glucose (17) one can readily postulate entrance into the cell, enzymic mutarotation, and exit into the blood stream all in one passage through the liver. The hind limb, however, contains primarily skeletal muscle, a tissue less permeable to glucose. In the experiments of Cahill et al. (17) no diffusion of glucose into muscle cells was demonstrated in the same experiments in which the glucose-U4 in liver tissue water was equal to that in plasma water. This suggests that the mutarotation taking place in muscle must be extracellular. Thus, it must be nonenzymic or muscle mutarotase must be in an extracellular site. Williamson and DiPietro (18) have presented evidence for enzymic activity in an extracellular site in mammalian heart muscle and perhaps this is true of liver mutarotase as well. Another way in which the liver might be reestablishing anomer equilibrium in the blood stream might be by an exchange mechanism. The glucose entering the liver not at equilibrium might be exchanged for equilibrium glucose which is released into the hepatic veins. Since the two curves in Fig. 5 are practically superimposable, such an exchange would have to be on a molecule for molecule basis. In addition, one might expect there to be an increase in the glucose-C4 content of liver over that in plasma in the experiments described by Cahill et al. (17) and this did not occur. In some experiments in Table 2 (dogs 111, 112, and I 17) small excesses of cu-n-glucose did get through the liver and appeared in the peripheral circulation coincident with the initial rise in total glucose. If this represented saturation of an exchange mechanism it might be expected to occur at a higher glucose level later in the infusion. The same argument would apply to saturation of an extracellular enzyme. In any case, experiments in which labeled glucose was infused into the portal vein and the specific activities of portal and hepatic vein blood determined during the initial phase of the infusion would determine whether such an exchange were actually taking place. The complete return to anomeric equilibrium of glucose infused into the femoral vein by the liver is consistent with its high mutarotase content. The other regions of the body vary tenfold in mutarotase content

78

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BIBER,

ASD

HUFFIh-ES

and yet do not differ significantly in their ability to reestablish anomeric equilibrium. The simplest explanation lies in the uneven distributon of mutarotase in these regions where the activity concentration may surpass even that of liver. The renal cortex, for example, contains 0.5 unit/gm of wet weight. The upper small intestine contains higher activity concentration than the stomach or large intestines (unpublished data). Thus, all the blood in the effluent veins has not passed through areas of equal mutarotase activity. As already noted, the permeability of various cells to glucose and the exact location of the enzyme in, on, or about the cell might be expected to influence enzymic mutarotation in these experiments. Since no measurements of blood flows were made in these experiments, no precise comparisons of the contributions of the various regions to mutarotation in terms of mg glucose mutarotated/gm of tissue/minute can be made. In view of the known high blood flows through the liver, the marked contribution of that organ to mutarotation is evident. The fact that anomer equilibration in the blood stream of the intact dog is more rapid than net glucose outflow from the bloodstream might be interpreted as evidence against a rate limiting role of mutarotase (8) in glucose transport and metabolism. It must be emphasized, however, that these experiments do not establish a relationship between in vivo mutarotation and mutarotase and further, it is possible that mutarotation could still be the rate limiting step in some regions of the body and not in others. Figure 7 shows an abrupt rise in the blood glucose a short while after the phlorizin infusion rate was increased. Similar changes have been noted after the administration of phlorizin (19, 20) but renal blood vessel ligation was usually necessary before they became apparent. This rise could not be accounted for on the basis of a slight fall in glucose excretion which did take place between periods 10 and 11. Stettin et al. (21) studying the oxidation of glucose-C’4 to C?“O, in normal and phloridzinized rats found a decreased conversion in the latter group. The authors attributed this to the decreased blood glucose levels, but a decrease in glucose oxidation secondary to a decrease in glucose transport into cells might be an equally plausible explanation. Although it is known that the use of creatinine for the measurement of glomerular filtration rate in the male dog gives somewhat erroneous results (22)) its use in the above experiments should not alter the conclusions. Absolute values are of minor importance in these experiments where the relative values obtained for the different anomers of glucose should be indicative of differences in the way these anomers are transported by the kidney. No such differences were observed.

MUTAROTASE

AKD

GLUCOSE

ANOh’lERS

7’1I

The normal mammalian kidney is rich in mutarotase and thus one might assume either anomer would be brought to equilibrium prior to being transported. Two pieces of evidence tend of discount this hypothesis. First, in the phloridzinized dog the anomer infused in excess can be found in the urine in excess (unpublished data). This means that glucose in the lumen of the renal tubule is not yet brought to equilibrium. Secondly, in the phloridzinized kidney in which glucose reabsorption is clearly impaired, the two anomers are similarly cleared. If mutarotase plays a role in glucose transport in the kidney, it is not apparent from tllcye experiments and does not occur by the selective transport of either the CY-or /3-n-glucose anomer. PART

II MATERIALS

AND

METHODS

Measurements of tissue mutarotase activity were done as previously described (14). When these experiments were initiated quantitation of the mutarotase activity in terms of unitage was not feasible. For this reason all analyses for a single experiment were completed at the same time. Subsequently a means of quantitation has been worked out, but in order that the data be consistent, they are presented, for the most part, as percentages of control values rather than as units of mutarotnse activity. Rats were maintained on Purina Chex for rats, mice, and hamsters and were allowed access to food and water throughout the experiments. Injections of the nephrotoxins were given subcutaneously. At the time of sacrifice 50 mg/kg sodium pentobarbital was administered intraperitoneally. The abdomen was opened with a midline incision and the animal was exsanguinated from a needle placed in the abdominal aorta or vena cava. Blood was collected, allowed to clot and the serum used for glucose determinations. The kidneys were removed, trimmed, weighed, and divided into three sections. One pole was kept in a commercial deep freeze for mutarotase assay, the other pole was used to determine wet and dry weight, and the center section was placed in a formalin solution for the preparation of microscopic slides. Tissues other than kidney were collected as noted in the text. In certain experiments urine was collected by placing the rat in a metabolism cage in which food spillage and urine contamination were minimized by a series of baffles. In some cases urine was collected at autopsy from the bladder. Qualitative glucose tests were accomplished using Clinistix (Ames Co. Inc., Elkhart, Indiana). Urine glucose levels were determined by a modification of the nuto-

80

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AND

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BIBER,

AND

HUFFINES

mated glucose-oxidase method (23). Urines were acidified and treated with Lloyd’s reagent prior to analysis. Severity of histological damage to the kidneys of the potassium dichromate treated rats was graded from O-5 by one of the experimenters (W. H.) before he knew the dosage of nephrotoxin administered. RESULTS

Table 5 contains data from experiments utilizing three nephrotoxic salts at the doses recorded. All three salts produce a fall in total kidney mutarotase activity 72 hours after their administration. Neither potassium dichromate nor uranyl nitrate produce an inhibitory effect on rat kidney mutarotase activity when added to the enzyme in vitro at levels TABLE ANIMALS

Treatment

TREATED

group

Control Potassium dichromate 15 mg/kg Uranyl nitrate 15 mg/kg Mercuric chloride 11 mg/kg

WITH

5

THREE

NEPHROTOXIC

Number of animals 9 8 5 4

S.~LTS

Mutarotase activity yO of control group mean (range) 100 63 69 46

(99-101) (50-79) (66-74) (21-55)

as high as 0.5 mg/ml of enzyme preparation. Mercuric chloride does produce in vitro inhibition at doses as small as 10 pg/ml. It is quite unlikely that the concentration of the dichromate and uranyl salts in the kidney extracts assayed would be higher than 0.5 mg/ml. The EDTA added to the extractant was adequate to protect the enzyme from destruction from any mercuric chloride contacted in the extraction process or the subsequent assay procedure. As may be seen from the table, all three salts produce significant falls in kidney mutarotase activity levels. All the rats had glucose in their urine and histologic damage to their kidney tubules. The mercuric chloride treated animals were visibly the sickest and it is doubtful they would have survived this dose. Fig. 10 contains dose response data on the relationship between the quantity of potassium dichromate administered and the kidney mutarotase activity found 72 hours later. There is a clear relationship which is equally evident whether the data is expressed on kidney wet weight or dry weight basis. It is of interest to note that at the higher dose, the range of kidney mutarotase activity values markedly narrows suggesting that the activity which is influenced by dichromate treatment has all been

MUTAROTASE

Effect

of

Potassium

AND

a Single

GLUCOSE

Subcutaneous

Dtchromafe

Mufarotase

Activity Hours

Activity/

wet

weight

81

ANOMERS

on

ln/ecflon the

Found

Rat

of Kidney

Seventy-

two

Later _Actlvity

/dry

weight

1 ~

IO

Dose

of

Potassium

Dichromote

15

20

mg/kg

FIG. 10. Effect of a single subcutaneous injection of potassium dichromate on the rat kidney mutarotase activity found 72 hours later; kidney mutarotase activity vs. dose of potassium dichromate administered.

destroyed and that which remains is in a portion of the kidney which is remote from dichromate destruction. That the tubular damage is accompanied by a loss of enzyme may be seen in another way in Fig. 11 in which mutarotase activity may be seen to be present in the urine of rats treated with 15 mg/kg potassium dichromate three days prior to the urine collection. The tracing was made by continuously sampling from a G-25 Sephadex column. The urine was placed on the column and the first peak following :L nulrkcr is the larger molecule protein enzyme fraction followed by the smaller molecule glucose fraction. Known mutarotase activity is destroyed by heating to 60’ for five minutes. This phenomenon is used here as a control to be certain the mutarotational catalyst is indeed mutarotase. Urine from untreated rats was not found to contain either peak when placed on the column. Similar results were obtained with urine from uranium and mercury treated rats. Thus the three salts produce a condition in which the enzyme is lost to the urine. The mercuric chloride does not destroy all of the enzyme even though it is capable of irreversibly inhibiting mutarotase at concentrations over 10 pg/ml.

82

HILL,

Rat

COWART,

A

0 85 ml uftne

AND

HILL,

BIBER,

AND

Rat

HUFFIKES

A

0 70 ml u,,ne healed to 63” Iof 5 mlnute5

G -25 Sephadex Column Rafs given /5mg/&g Pofassium Dichromale 3 days prior to Urine Colecfion FIG. 11. Continuous monitoring of column effluent for mutarotase activity. The first spike in each tracing is a glucose marker which denotes the beginning of monitoring of the effluent. Absorbancy is on the ordinate.

As may be seen in Table 6 there is a general correlation between the urinary glucose concentration, the severity of histological damage, the mutarotase activity lost from the kidney, and the dose of potassium dichromate administered. There are specific instances, as for example rat 12, where the correlation does not seem to hold. Whether thcFe are owing to the limitations of our measurements or to the exact location in the tubule of the lesions is not known. If animals treated with dichromate are allowed to recover for 21 days, the mutarotase activity content returns toward the preinjection levels. This may be seen in Table 7. There was a concomitant loss of glycosuria as well as histological evidence of recovery. Table 8 contains data obtained from mutarotase assays of tissues other than kidney. It is apparent that the three nephrotoxins do not exert much of an influence on the tissue mutarotase activity of the tissues listed.

SO~UE EFFECTS

MCTAROTASE

AXD

OF DICHROMATE HISTOLOGY,

TABLE 6 INJECTION AND ENZYME

Treatment

Rat

Control

GLUCOSE

Urinary glucose mg/lOO ml

83

ANOMERS

ON 1i.k~ KIINEY CONTEXT

FUNCTION,

Estimate of severity of histological damage (O-51

8

Mutarotase activity/wet weight r. of control

3 4 5 6

5 1 4 5 5

1 0 1 1 1 1

104 106 l(12 94 !)2 10%

7 8 9 10

7 4 10 15

3 0 0 2

74 !)6 10X ss

15 mg/kg

11 12 13 14

2750 226 158 56

.5 3 4 5

76 6X 86 70

20 mg/kg

15 16 17 18

136 130 318 361

*5 5 5 5

40 44 36 3x

1 2

Potjassium

dichromate

10 mg/kg

I~IUNEY

MUTAROTASE

ACTIVITY

TABLE 7 3 DAYS AND 21 DAYS DICHROMATE

AFTER

I’OTASSITJ~

Mutarotase activity/ gram of wet, weight No.

Control” Sacrificed 15 mg/kg Kidney Kidney Nephrectomy Kidney Kidney a Data

animals used

Day 3 Potassium dichromate removed day 3 removed day 21 Controls removed day 3 removed day 21 expressed

as ye of csntrol

values.

No. kidneys assayed

Mean

(range)

2

4

100 (88-106)

5

5 5

67 (46-82) 86 (8%90,

5

5 5

93 (85-97) 92 (79-101)

84

HILL,

COWART,

ANL)

HILL,

BIUEIZ,

AXI)

RII~FISES

TTnits/gm of wet weight Hat number experiment 1 3/18

Treatment

group

Control

3 3/18 1 6/7 2 O/7 3 3/18 4 3118 5 6 7 S 9

S/18 $48 6/7 6/7 6/7

4 6/7 5 B/7 6 6/7

Potassium dichromate 16 mg/kg Uranyl nitrate 15 mdk

Mercuric chloride 11 w/k

Skeletal muscle

Live1

LlUl~

0.60 0.54

0 0:3 0.02

0.0” 0. 0:;

0.51

0.03

0 .o:!

0.11 0.09

0 Cl!)

0.50

I) Ga

IJ 0;:

0.10

0.56 0.50

0.0:: 0 03

O.(G 0.04

0.10

o.,‘,:! 0 5s 0.56 0 63 0 tit

0.03 0 .o:: 0 0:; 0 .0:-i 0 04

0 04 0.04 0.0.5 0.0” 0 0:i

0.09 0 OS 0. (I!) 0 OR 0.11

0.3” 4 0.51 0 2s

0.03 0 .03 0. 03

0.0’2 0.02 0, OS

0 .09 0.09 0 O!l

0.0s

While there is suggestive evidence of lowering of the liver mutarotase activity by mercury bichloride, this is not surprising in that levels found in mouse liver four days after a single injection are second only to the kidney (24). A possible elevation in the skeletal muscle of the uranium treated rats is interesting in veiw of the reported finding of elevated alkaline phosphatase in the kidneys following treatment with this s;dt (13). More experiments would be necessary to establish this point. DISCUSSION

Oliver et al. (25) using a technique of microdissection showed that potassium dichromate, mercuric chloride, and uranyl nitrate produced lesions limited to the proximal convolution of the renal tubule of dogs and rabbits. This finding coupled with the results presented here suggests that the majority of mutarotase activity in the rat kidney is associated with the proximal tubule. The micropuncture studies of Walker et al. (26) locate glucose reabsorption in the proximal tubule. While the present studies do not indicate whether mutarotase activity is necessary for glucose reabsorption, they do show that in specific situations in which there is proximal tubular damage and glycosuria, there is also a decrearc in mutarotase activity.

MUTAROTASE

AND

GLUCOSE

AXOMERS

83

SUMMARY

The mutarotation of exogenous glucose by various body regions of the anesthetized dog has been studied. The liver, an organ rich in mutarotase activity was found to produce the greatest changes in relative anomer content of the blood flowing through it. The reabsorption of glucose by the kidney of the anesthetized dog is not altered by infusions containing an excess of either (Y or P-n-glucose over that found at thermodyamic equilibrium. Phlorizin, in doses which inhibit glucose reabsorption does not inhibit the reabsorption of cy- or /3n-glucose to a different degree. Rats treated with nephrotoxic doses of potassium dichromate, mercuric chloride, and uranyl nitrate were found to have glycosuria, histological damage to the renal tubule, and a decreased renal mutarotase activity. Mutarotase activity was found in the urine 72 hours after subcutaneous administration of these nephrotoxins. Histological recovery was accompanied by a loss of glycosuria and an increase in the enzyme activity. Heart muscle, skeletal muscle, and Iung tissue were not affected by the treatment though liver enzyme activity may have been lowered by treatment with mercuric chloride. ACKNOWLEDGMENTS This work has been supported by a grant (No. AM 09033) from the National Institute for Arthritis and Metabolic Diseases of the National Institutes of Health, U. S. Public Health Service. The authors would likr t,o thank Dr. Albert S. Keston of the Institute for Medical Research and Studies for his constructive criticism, help and encouragement throughout many phases of this work.

1. HILL, J. B., 2. LUNDSGAARD, 3. 4. 5. 6.

7. 8. 9.

10. 11. 12. 13.

14. 15.

J. AppZ. Physiol. 20, 749 (1965). C., AND HOLB@LL, S. A., J. Biol. Chem. 62, 453 (1924-1925); J. Biol. Chem. 65, 343 (1925). BENTLEY, R., AND NEUBERGER, A., Biochem. J. 45, 584 (1949). KEILIN, D., AND &TREE, E. F., B&hem. J. 50, 341 (1952). KESTON, A. S., Science 120,355 (1954). BA~EY, J. M., FISHMAN, P. H., AND PENTCHEV, P. G., Science 152, 1270 (1966). WALLENFELS, K., AND HERRMANN, K., B&hem. Zeit. 343,294 (1956). KESTON, A. S., J. Biol. Chem. 239,324l (1964). STUECICF,R, H. J., AND KORKES, S., J. Biol. Chem. 196,769 (1952). C~LOWICK, S. P., AND GOLDBERG, E. B., Bull. Res. Council Israel llA4, 373 (1963). JOEGENSEN, B. B., AND JORQEN~EN, 0. B., Acto Chem. Scund. 20, 1437 (1966). FEIANK, E., Arch. Ezptl. Puthol. Pharmakol. 72,387 (1913). HEPLER, 0. E., AND SIMONDS, J. P., Arch. Pathol. 41,42 (1946). HILL, J. B., AND COWART, D. S., Anal. B&hem. 16,327 (1966). CHASSON, A. L., GRADY, H. J., AND STANLEY, M. A., Am. J. Clin. Pathol. 35, 83 (1961).

86

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AND

HILL,

BIBER,

AND

HUFFIiYES

16. REICHARD, G. A., FRIEDMAN, B., MAASS, A. R., AND WEINHOVSE, S., J. Biol. Chem. 230, 387 (1958). 17. CAHILL, G. F., JR., ASHMORE, J., EARLE, A. S., AND ZOTTU, S., Am. J. Physiol. 192(3), 491 (1958). 18. WILLIAMSON, J. R., AND DIPIETRO, D. L., Biochem. J. 95,226 (1965). 19. UNDERHILL, F. P., J. BioZ. Chem. 13, 15 (1912-13). 20. CSONKA, F. A., J. Biol. Chem. 26, 93 (1926). 21. STF,TTIN, D., JR., WELT, I. D., INGLE, D. J., AND MORLEY, E. H., J. Biol. Chem. 192, 817 (1951). 22. O’CONNELL, J. M. B., ROMEO, J. A., AND MVDCE, G. H., Am. J. Physiol. 203, 985 (1962). 23. HILL, J. B., AND KESSLER, G., J. Lab. Clin. Med. 57,970 (1961). 24. BERLIN, M., AND ULLBERG, S., Arch. Environ. Health. 6, 589 (1963). 25. OLIVER, J., MACDOWELL, M., AND TRACY, A., J. Clin. Invest. 12, 1305 (1951). 26. WALKER, A. M., BOTT, P. A., OLIVER, J., AND MACDOWELL, M. C., Am. J. Physiol. 134, 580 (1941).