Metabolic changes during spermatogenesis and thoracic tissue maturation in Drosophila hydei

DEVELOPMENTAL

BIOLOGY

Metabolic

28, 390-406 (1972)

Changes

during

Spermatogenesis

Tissue Maturation

in Drosophi’la

and Thoracic hydei’

B. W. GEER, D. V. MARTENSEN,~ B. C. DOWNING,~ AND G. S. MUZYKA Department of Biology, Knox College, Galesburg, Illinois 61401 Accepted

February

11, 1972

In oitro enzyme studies indicate that gametes depend upon carbohydrate oxidation by way of the pentose shunt cycle, glycolysis, and the Krebs cycle for energy early in spermatogenesis in Drosophila hydei. Enzymes associated with amino acid metabolism and fatty acid synthesis are also high in activity at these developmental stages. In late spermiogenesis gametic glycolytic and Krebs cycle enzyme activities increase greatly. Both carbohydrates and amino acids are likely energy sources for mature spermatozoa. Low lactate dehydrogenase and @-hydroxyacyl dehydrogenase activities suggest that gametes at all developmental stages have low capacities for lactic acid formation and fatty acid oxidation. Thoracic tissue development differs fundamentally from testis development in that the a-glycerophosphate dehydrogenase-oxidase cycle becomes highly developed in thoracic tissue, whereas the low activity of cY-glycerophosphate dehydrogenase indicates that it is a minor factor in gamete metabolism. Furthermore, NADPisocitrate dehydrogenase and malic enzyme become increasingly less active as thoracic tissue matures, with malate dehydrogenase and NAD-isocitrate dehydrogenase becoming more active. The NADP-dependent enzymes are very active at all stages of spermatozoan development. Glycolytic and Krebs cycle enzyme activities are much higher in thoracic tissue at all maturation stages than in the mature testis.

sperm and spermatids in different ways (Sobels, 1965). Recent cytogenetic and biochemical studies suggest the means by which metabolic changes in developing gametes may be facilitated. Although RNA synthesis is premeiotic in developing sperm, Beermann, Hess, and Meyer (Beermann, 1965, 1967; Hess, 1966, 1967; Hess and Meyer, 1968) hypothesized that “lampbrush-like” configurations in the primary spermatocyte of Drosophila hydei represent sites of long-lived RNA” synthesis. According to this hypothesis, long-lived messenger RNA for the synthesis of a variety of proteins during spermiogenesis may be

INTRODUCTION

It is apparent from genetic studies that the morphological changes that occur during spermatogenesis in Drosophila are accompanied by changes in metabolism. Cells representing different stages of spermatogenesis have different radiosensitivities which seemingly reflect metabolic differences in the developing gametes (Baker and Von Halle, 1953; Sobels and Tates, 1961; Liming, 1961; Oster, 1961; Lefevre and Jonsson, 1964; Traut, 1964; Trout, 1964; Sobels, 1965; Shiomi, 1967). Oxidative respiration appears to differ in spermatids and mature sperm since alteration of respiration by inhibitors and changes in oxygen tension affect the frequencies of maturation induced in mature

‘The abbreviations used are: CoA, coenzyme A; DNA, deoxyribonucleic acid; EDTA, ethylenediaminetetraacetic acid; NADH, reduced nicotinamide adenine dinucleotide; NAD+, oxidized nicotinamide adenine dinucleotide; NADPH, reduced nicotinamide adenine dinucleotide phosphate; NADP+ oxidized nicotinamide adenine dinucleotide phosphate; RNA, ribonucleic acid; Tris, trisfhydroxymethyl)aminomethane.

‘This research was supported by National Science Foundation Grant GB-13393. 2Present address: Department of Genetics, University of California, Davis, California. 3Present address: Department of Zoology, Utah State University, Logan, Utah. 390 Copyright

0 1972 by Academic

Press. Inc.

GEER et al.

Testis

and Thoracic

formed by the transcription of a number of genes in the primary spermatocyte genome. Sperm development during spermiogenesis is ostensibly regulated by the long-lived RNA. As evidence for the hypothesis, Hennig (1968) has found RNA in the testis of D. hydei complementary to Y-chromosome DNA. Both chromosomal and enzymatic protein appear to be synthesized during spermiogenesis in Drosophila melanogaster. Brink (1968) found that protein synthesis occurs in early and elongating spermatids, a developmental period which would include the histone transformation observed by Das et al. (1964). Furthermore, carnitine acetyltransferase, an enzyme associated with energy-yielding metabolism, increases greatly in activity during spermiogenesis (Geer and Newburgh, 1970). In this paper we report changes in testicular enzyme activities that coincide with sexual maturation in D. hydei males. To relate these observations we propose the following model for changes in the energy-yielding metabolism of developing gametes during spermatogenesis in Drosophila. Division of the spermatogonia results in a number of spermatocytes that increase greatly in size prior to meiosis. Growth of the primary spermatocyte is accompanied by the synthesis of materials to be used during spermiogenesis. This synthesis is supported by carbohydrate oxidation by way of the pentose shunt cycle, glycolysis, and the Krebs cycle, and to a lesser extent by amino acid and fatty acid oxidations. As the gametes near maturation, presumably as late stage spermatids, their capacity to degrade carbohydrate and amino acids increases 2- to 4fold as compared to earlier stages of spermatogenesis, but more strikingly the capacity to degrade carbohydrate anaerobically to pyruvate and then to alanine or some alanine derivative increases. Acetylcarnitine may also form as the result of glycolysis. Both alanine and acetylcarnitine, once formed, may be metabolized

Tissue

Maturation

391

readily under aerobic conditions by conversion to acetyl-CoA and subsequent degradation by the Krebs cycle reactions. The Krebs cycle capabilities of mature Drosophila spermatozoa are greater than at earlier stages and are sufficient for mature spermatozoa to acquire energy by aerobic degradation of both carbohydrate and amino acids in addition to anaerobic degradation of carbohydrate. Drosophila spermatozoa are not capable of forming lactic acid as a major end product of glycolysis or oxidizing fatty acids at a rapid rate. In addition to the experimentation that led us to formulate this model, we report observations of metabolic changes that accompany thoracic tissue maturation. We conclude that both testis and thoracic tissues possess unique developmental metabolic patterns. METHODS AND MATERIALS

Animals. The D. hydei wild-type strain employed throughout the experimental work was established originally by Dr. H. D. Berendes at the Genetisch Laboratorium, Rijksuniversiteit te Leiden, Leiden, Netherlands and had been maintained under mass culture conditions for several years in the Department of Genetics, University of California at Davis and in this laboratory before its use in this study. Flies were raised and maintained on a cornmeal-yeast-sucrose-agar medium (Geer and Newburgh, 1970) under uncrowded conditions at 23.8”C with a lo-hr day-14 hr night. Adults were used for experimentation within 12 hr after eclosion, or were aged in half-pint cultures for the time periods given in the Results section. Homogenate preparation. Thoracic tissues were obtained by removing the head and abdomen from the thoraces of adult males, and testes were acquired by dissecting the testes of adult males and freeing them from adjoining tissues. Although the thoracic tissue may be re-

392

DEVELOPMENTALBIOLOGY

garded as consisting mostly of flight muscle, small amounts of leg and digestive tract tissues were also present in the preparations. Dissections were routinely performed in a drop of Drosophila Ringer solution (Ephrussi and Beadle, 1936) on a depression slide using jeweler’s forceps. The tissues were washed in a second drop of Drosophila Ringer solution before transfer to the homogenizing solution. Fifteen thoraces were homogenized in 200 ~1 of homogenizing solution using a ground-glass homogenizer. Forty pairs of testes from newly eclosed males or 30 pairs of testes from older males were homogenized in 200 ~1 of homogenizing solution. Homogenates were allowed to stand for 30 min at 4”C, then centrifuged for 20 min at 200 g at 4°C to remove gross debris. No further attempt was made to fractionate the components of the tissue homogenates, consequently the enzyme determinations reported in this study are total enzyme activities. To assess enzyme activities in different testis regions, the testes of lo-day-old males were dissected into three parts. The posterior region, extending from the point of attachment of the testis to the ejaculatory bulb to the middle region, was approximately 0.7 cm in length. The middle region, immediately anterior to the posterior region, was about 1.4 cm in length. The anterior region, extending from the middle region to the anterior tip of the testis, was about 0.4 cm in length. Routinely, 35 pairs of testes from lo-day-old males were dissected into the three regions in Drosophila Ringer solution, transferred to 100 ~1 of homogenizing solution, and homogenized with a ground-glass homogenizer. The homogenates of the testis regions were then prepared for enzyme analysis by the same procedures described for whole testis and thoracic tissues. Gametic contents of different regions in the immature and mature testes of D. hydei were estimated by direct measure-

VOLUME 28. 1972

ment with a phase-contrast microscope equipped with an eyepiece micrometer. Areas of the testis filled with spermatogonia, spermatocytes, and spermatids were quantified, and the proportion of the testis occupied by each stage was calculated by dividing the length of the testis filled with each stage by the total testis length. The testis was assumed to be uniform in diameter along its entire length. Calculations are based upon observation of five pairs of testes for each age group. Lactate dehydrogenase, aldolase, acetylCoA carboxylase, malate dehydrogenase, glucose-6-phosphate isomerase, pyruvate kinase, a-glycerophosphate dehydrogenase, and triosephosphate isomerase determinations employed homogenates of tissues prepared with 0.15 M KCl-0.02 M KHCOB, pH 7.8 (Ward and Schofield, 1967). Hexokinase was assayed using preparations made with 0.15 M KCl-0.05 M KHCO,-0.006 M Na,H,EDTA-0.005 M MgC12, pH 8.2 (Ward and Schofield, 1967). Tissues for L-aspartate aminotransferase and malic enzyme determinations were homogenized in 0.15 M KCl-0.05 M KHCO,-0.01 M 2-mercaptoethanol, pH 8.2. Phosphoglucomutase, glucose-6-phosphate dehydrogenase, @-hydroxyacyl dehydrogenase, citrate synthase, and 6phosphogluconate dehydrogenase were assayed using tissue preparations made with 0.1 M KH,PO,-K,HPO,, pH 7.3 with 0.002 M Na,H,EDTA (Beenakkers, 1969). Phosphoglyceromutase and enolase, the pyruvate dehydrogenase complex, Lalanine aminotransferase, phosphofructokinase, 3-phosphoglycerate kinase, glyceraldehyde-3-phosphate dehydrogenase, NAD-isocitrate dehydrogenase, NADPisocitrate dehydrogenase, and carnitine acetyltransferase determinations were made using tissue homogenates prepared with 0.1 M KH,PO,-K,HPO,, pH 7.3 with 0.002 M Na,H,EDTA and 0.01 M 2mercaptoethanol. Spectrophotometric enzyme assays. The assays were performed by following the

CTEER

t’t

al.

1 ems ana ’ Thoracic

oxidation of NADH or NADPH or the reduction of NAD+ or NADP+ in the reaction mixture at 30°C by measuring the change in extinction at 340 nm using a DU spectrophotometer Beckman equipped with a Gilford automatic cuvette positioner, Haacke constant temperature circulator, and Honeywell recorder. Assays were performed using 2-20 pg of homogenate protein in a total reaction volume of 0.5 ml. Each enzyme was assayed 3 to 6 times using different amounts of enzyme preparation to be certain that an increase in enzyme preparation content was accompanied by a corresponding increase in enzyme activity. Specific enzyme activities are given as the number of nanomoles of cofactor oxidized or reduced per milligram of homogenate protein per minute. Corrections for nonspecific changes in extinction were made when calculating enzyme activities by subtracting the rate of oxidation or reduction of cofactor during a 5-min preincubation period without substrate from that observed with the substrate. Reactions were started by the addition of substrate. The pyruvate dehydrogenase complex was assayed by supplying pyruvate and CoA-SH in the reaction mixture along with L-malate, malate dehydrogenase, and citrate synthase. The formation of acetylCoA from pyruvate and CoA-SH was assessed by following the condensation of acetyl-CoA with oxaloacetate to yield citrate. For each molecule of acetyl-CoA formed through the action of the pyruvate dehydrogenase complex in the reaction mixture, two molecules of NADH were formed. The reaction mixture contained Tris HCl buffer, pH 8, 100 pmoles; Na ,H 2EDTA, 2.5 pmoles; MgC12, 3.5 ~moles; thiamine pyrophosphate, 0.18 pmole; sodium pyruvate, 1.0 gmole; CoA-SH, 0.15 pmole; L-malate, 15.1 pmoles; NAD+, 2.5 pmoles; 2-mercaptoethanol, 3.5 pmoles; malate dehydrogenase, 20 units; and citrate synthase, 0.9 unit. P-Hydroxyacyl dehydrogenase (EC

Tissue

Maturation

393

1.1.1.35) was measured by following the oxidation of NADH in a reaction mixture containing KH ,PO,-K ,HPO d buffer, pH 6.8, 60 Fmoles; Na,H,EDTA, 5.0 pmoles; NADH, 0.13 pmole; and S-acetoacetylCoA, 0.05 hmole. This reaction mixture is a modification of that used by Decker (1965) for the determination of S-acetoacetyl-CoA. Other enzymes were assayed according to methods given in the references of Table 1. The acetyl-CoA carboxylase assay procedure was modified by reducing the concentration of acetylphosphate to one-third that employed by Matsuhashi et al. (1964) to obtain maximum activity for the enzyme in Drosophila enzyme preparations. Isotopic enzyme assays. Carnitine acetyltransferase (EC 2.3.1.7) was assayed by the isotopic method of McCaman et al. (1966). Protein determination. Protein was measured by the method of Lowry et al. (1951). Crystalline bovine albumin was used as the standard. Sex maturation tests. Two tests were performed to assess the degree of maturation of gametes in the testes of maturing D. hydei males, sperm motility and fertility tests. To determine the presence or absence of motile sperm in the testis, testes from lo-25 males of a given age were teased apart in Drosophila Ringer solution. Sperm capable of motility became active when exposed to Drosophila Ringer solution and were easily detected by light microscope observations. Fertility tests were performed by housing lo-25 males of each age group individually with three sexually mature virgin females for 24 hr. If any of the three females subsequently reproduced, the male was judged to be fertile. To correlate sexual maturation with testis development, two additional observations were made. Testis lengths for adult males were assessed by measurement with an eyepiece micrometer

394

DEVELOPMENTAL

BIOLOGY

VOLUME

TABLE METHODS

USED FOR ENZYME

Name of enzyme Glycolysis Hexokinase Phosphoglucomutase Glucose 6-phosphate isomerase Phosphofructokinase Aldolase o-Glycerophosphate dehydrogenase Triosephosphate isomerase Glyceraldehyde 3-phosphate dehydrogenase 3-Phosphoglycerate kinase Phosphoglyceromutase and enolase” Pyruvate kinase Lactate dehydrogenase Krebs cycle Citrate synthase NAD-isocitrate dehydrogenase Malate dehydrogenase NADP-dependent Glucose B-phosphate dehydrogenase 6-Phosphogluconate dehydrogenase NADP-isocitrate dehydrogenase Malic enzyme Amino acid metabolism L-Alanine aminotransferase L-Aspartate aminotransferase Fatty acid metabolism Acetyl-CoA carboxylase 0 The reaction

mixture

Acnvm

28, 1972

1

DETERMINATIONS

IN THE CURRENT

STUDY

Systematic EC number

Reference

2.7.1.1 2.7.5.1 5.3.1.9 2.7.1.11 4.1.2.b 1.1.1.8 5.3.1.1 1.2.1.12

DiPietro and Weinhouse (1960) Najjar (1946) Shonk and Boxer (1964) Underwood and Newsholme (1965) Beizenherz (1955) Beenakkers (1969) Shonk and Boxer (1964) Ward and Schofield (1967)

2.7.2.3 2.7.5.3 4.2.1.11 2.7.1.40 1.1.1.27

Ward and Schofield Beames (1963)

4.1.3.7 1.1.1.41 1.1.1.37

Beenakkers (1969) Goebell and Klingenberg (1964) Bergmeyer and Bemt (1965a)

1.1.1.49 1.1.1.43 1.1.1.42 1.1.1.40

Glock and McLean (1953) Horie (1967) Komberg (195513) Ochoa (1955)

2.6.1.2 2.6.1.1

Mattenheimer (1970) Bergmeyer and Bemt (1965b)

6.4.1.2

Matsuhashi

measured the combined activities

mounted in a dissecting microscope. Testes from five males of each age group were examined and an average testis length was determined. To assess the testis protein content for the males of a given age group, five pairs of testes were homogenized in 200 ~1 of Drosophila Ringer solution and three different quantities of homogenate assayed for protein content by the Lowry et al. method (1951). Reagents and enzymes. Acetyl-l-14CCoA was purchased from the New England Nuclear Corporation. Acetyl-CoA, DL-carnitine-HCl, D-glucose, D-fructose, Lcysteine, L-aspartic acid, L-glutamic acid, and L-alanine were acquired from Nutritional Biochemicals Corporation. All other

(1967)

Tietz and Ochoa (1958) Komberg (1955a)

et al., (1964)

of the two enzymes.

organic reagents and enzymes were the highest grade available from the Sigma Chemical Company. Inorganic chemicals were reagent quality. RESULTS

Sexual Maturation Examination of D. hydei males of different ages showed that males become sexually mature B-10 days after eclosion. At 8 days of age 14% of the males possessed motile sperm and by 9 days of age all the males had motile sperm. In a separate experiment 10% of the s-day-old males, 78% of the g-day-old males, and 100% of the lo-day-old males successfully

GEER et al.

Testis

and Z’horacic

inseminated mature virgin females. The slight differences between the two tests may indicate that sperm become motile in the testis of D. hydei shortly before the males become fertile. Testis length and testis protein content measurements indicated that sexual maturation was delayed until testis development was completed. Testis length (Fig. 1) increased steadily from a length of 0.81 cm at eclosion to 2.45 cm at 8 days of age, the time that motile sperm first appeared in the testis. Only a slight increase in testis length was detected in males after 8 days of adult life. Testis protein content (Fig. 2) also increased from 9.6 pg of pro-

FIG. 1. The average testis length of testes from Drosophila hydei males of different adult ages.

Tissue

Muturation

395

tein per pair of testes at eclosion to 40.2 pg per pair of testes at 8 days of age. The increase in testis size during the adult maturation period reflected a change in gametic content. Whereas the region of the testis occupied by the spermatocytes and spermatogonia changed little in volume during maturation, the testicular volume of gametes undergoing spermiogenesis increased greatly. At eclosion the gametic content in terms of volume was approximately 37% spermatogonia and spermatocytes, and 63% early spermatids. At sexual maturation spermatogonia and spermatocytes constituted about 11% and spermatids and mature sperm 89% of the total testis gametic content. The spermatogonial and spermatocyte contents were lumped together because of the difficulty in quantifying the spermatogonium content. In the region of the testis occupied by spermatocytes and spermatogonia the ratio of spermatocyte volume to spermatogonium volume was at least 15: 1. In both immature and mature males the gametes comprised about 90% of the total volume of the testis. In any event, the increase in testis length and protein content during sexual maturation was due primarily to the development of late stage gametes. Testis Metabolism

AO-

30-

1

2

3

4

5

6

7

8

9

AGE FIG. 2. The average testicular protein content Drosophila hydei males of different adult ages.

of

Glycolytic enzymes. The activities of the enzymes of the glycolytic pathway (Table 2) indicate that glycolysis is more active in the testis of sexually mature males than in immature males. With the exception of glyceraldehyde 3-phosphate dehydrogenase, whose activity in the testes of both young and mature males was high and probably nonlimiting, all the glycolytic enzymes exhibited higher specific activities in lo-day-old male testes than in the testes of newly eclosed males. Carbohydrate degradation appeared to be primarily by way of the Krebs cycle in the mature testis since enzymes representing

396

DEVELOPMENTAL BIOLOGY

VOLUME 28, 1972

TABLE 2 GLYCOLYTIC ENZYME ACTIVITIES IN l- AND lo-DAY-OLD

MALE Drosophila

Testis

hydei Thorax

Enzyme Phosphoglucomutase Hexokinase fructose glucose Glucose &phosphate isomerase Aldolase Phosphofructokinase Triosephosphate isomerase Glyceraldehyde 3-phosphate dehydrogenase 3-Phosphoglycerate kinase Phosphoglyceromutase and enolase Pyruvate kinase Lactate dehydrogenase a-Glycerophosphate dehydrogenase a The mean activity

1 Day

10 Day

1 Day

10 Day

16.6 i 2.4”

21.8 + 3.8

21.1 * 0.7

60.0 i 3.5

13.2 f 1.3 28.1 + 4.1 418.1 * 67.8

47.2 i 6.7 38.8 + 2.6 568.2 zt 96.0

56.6 * 2.8 70.4 l 7.5 1082.8 zt 116.4

89.1 iz 11.8 88.9 i 6.6 1543.8 * 123.4

4.6 2.4 3624.7 406.6

* + + zt

2.1 1.1 420.0 34.2

44.4 9.7 4036.2 392.7

zt * zt +

2.3 1.4 707.4 23.4

267.6 79.5 18499.1 2583.5

i * * *

29.7 19.1 5534.6 152.1

576.7 187.5 29278.4 2870.1

+ 35.3 i 16.4 i 5711.6 zsz102.3

52.6 * 8.3 30.9 * 6.1

86.0 * 12.1 65.5 zt 4.6

452.0 * 70.1 231.9 * 54.5

778.8 * 30.5 396.8 f 52.2

14.3 i 2.7 31.7 zt 6.2 9.3 zt 0.8

75.6 f 3.2 18.5 zt 4.8 4.4 i 0.6

153.0 * 19.8 9.6 * 5.7 184.5 * 31.8

561.4 + 86.4 4.0 i 2.4 361.0 i 44.4

+ the standard

deviation.

,

TABLE 3 KREBS CYCLE ENZYME ACTIVITIES IN l- AND lo-DAY OLD MALE Drosophila Testis

hydei Thorax

Enzyme 1 Day Pyruvate dehydrogenase complex Citrate synthase NAD-isocitrate dehydrogenase Malate dehydrogenase o The mean activity

+ the standard

232.3 94.6 4.5 568.2

zt 28.4O + 8.4 +c 0.6 zt 147.0

10 Day 325.4 124.7 9.0 599.2

+ + i +

29.6 4.2 0.6 66.7

1 Day 521.2 292.4 7.6 717.3

zt 101.9 zt 73.2 * 1.8 z+z158.2

10 Day 1011.2 653.7 30.5 1514.1

zt zt * +

178.2 75.6 7.5 48.8

deviation.

other routes of degradation, lactate dehydrogenase and a-glycerophosphate dehydrogenase, declined in activity in the testis from the time of eclosion to sexual maturation. On the other hand, pyruvate dehydrogenase complex activity, active in the formation of acetyl-CoA for Krebs cycle consumption, increased by about one-third during the maturation period (Table 3). Lactate dehydrogenase activity, though diminished, appeared high enough in the mature testis to be a moderate factor in the metabolism of certain cells, but a-glycerophosphate dehydrogenase activity was so low that it is highly un-

likely that it is a major factor in the metabolism of the mature testis. Krebs cycle enzymes. Activities of the Krebs cycle enzymes evidenced a greater role for Krebs cycle activity in the mature testis than in the immature testis (Table 3). Citrate synthase and malate dehydrogenase were both relatively active in the testis of young males, and increased only slightly in activity in the mature testis. NAD-isocitrate dehydrogenase, which may have been the limiting Krebs cycle enzyme, was twice as active in the mature testis as in the immature testis. NADP-dependent enzymes. In contrast

GEER

et

al.

Testis

and Thoracic

to the glycolytic and Krebs cycle enzymes, the activities of NADP-dependent enzymes (Table 4) were high in both sexually immature and mature testes. The activities of glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase indicated that the pentose shunt cycle is a factor in carbohydrate metabolism in immature and mature testes, but is of relatively greater importance in the immature testis since glycolytic enzyme activities increased during maturation while the pentose shunt cycle enzyme activities remained essentially the same. That malic enzyme and NADP-isocitrate dehydrogenase activities were also high in both mature and immature testes suggested that NADPH generation by these enzymes and the pentose shunt cycle enzymes may be important in testicular metabolism. Amino acid and fatty acid metabolism enzymes. Carnitine acetyltransferase activity increased strikingly in the testis during sexual maturation (Table 5);

Tissue

397

Maturation

whereas P-hydroxyacyl dehydrogenase, a fatty acid oxidation enzyme, declined markedly and acetyl-CoA carboxylase, a fatty acid synthesis enzyme, dropped slightly in activity. The two amino acid metabolism enzymes tested had high activities; L-alanine aminotransferase more than doubled in activity in the testis during maturation while L-aspartate aminotransferase was high in activity in both immature and mature testes. Thus, carnitine acetyltransferase activity was directly related to pyruvate metabolism, paralleling the activities of pyruvate kinase, Lalanine aminotransferase, and the pyruvate dehydrogenase complex. Carnitine acetyltransferase activity was not directly correlated to the fatty acid metabolism enzyme activities. Intermediate developmental stages. Several enzymes were examined in whole testes at intermediate stages of maturity (Table 6). The glycolytic enzymes, hexokinase and 3-phosphoglycerate kinase, increased markedly in activity in the 4 days

TABLE 4 NADP-DEPENDENT ENZYME ACTIVITIES IN l- AND 10.DAY-OLD MALE Drosophila Testis

hydei Thorax

Enzyme 1 Day Glucose 6-phosphate dehydrogenase 6-Phosphogluconate dehydrogenase Malic enzyme NADP-isocitrate dehydrogenase a The mean activity

* the standard

45.1 12.4 220.7 105.9

i + * *

10 Day 42.9 9.8 219.2 103.0

4.6” 2.3 18.0 2.7

& * * *

2.3 1.9 39.0 7.2

1 Day 26.6 12.1 84.5 108.3

A * i +

10 Day 0.8 1.5 9.8 9.3

29.9 11.4 37.6 55.0

* 1.0 f 1.1 z+z6.3 + 7.6

deviation.

TABLE 5 FATTY ACID AND AMINO ACID METABOLISM ENZYME ACTIVITIES IN l- AND lo-DAY-OLD Drosophila hydei Testis

MALE

Thorax

Enzyme 1 Day Carnitine acetyltransferase ,!f-Hydroxyacyl dehydrogenase Acetyl-CoA carhoxylase L-Aspartate aminotransferase L-Alanine aminotransferase a The mean activity

51.8 55.9 10.2 311.1 188.9

* * * zt +

* the standard deviation.

4.7” 9.1 4.2 31.8 41.4

10 Day 70.2 22.9 8.1 362.9 422.7

* * * zt +

6.5 4.1 3.0 42.6 28.4

1 Day 47.9 73.6 3.5 187.2 516.0

* i * i +

5.6 2.2 1.0 12.0 46.8

10 Day 43.6 49.2 4.1 226.5 453.2

f * * * i

5.4 3.1 1.0 13.8 33.6

398

DEVELOPMENTAL BIOLOGY

VOLUME 28, 1972

TABLE 6 ENZYME ACTIVITIES AT DIFFERENT ACES IN MALE Drosphih

hydei

Age

Enzyme 1 Day

4 Days

6 Days

10 Days

Testis Hexokinase fructose glucose 3-Phosphoglycerate kinase Malate dehydrogenase Malic enzyme Carnitine acetyltransferase

13.2 21.7 52.6 485.0 220.7 45.6

+ f zt f f i

1.3” 1.5 8.3 56.2 18.0 4.7

16.0 23.6 46.9 99.8 235.9 100.3

zt f zt zt * *

2.8 3.6 7.6 7.8 59.0 7.7

19.7 37.7 46.6 212.9 228.8 80.0

* * + + zt

3.0 9.1 8.4 13.2 34.9 6.4

42.6 63.6 86.0 599.2 219.2 80.5

* f * + + *

14.4 10.2 35.1 78.0 8.0 4.7

104.7 129.6 778.8 1298.1 37.6 44.2

l

f 10.8 z+z9.0 f 12.1 zt 26.5 + 39.0 + 6.6

Thorax Hexokinase fructose glucose 3-Phosphoglycerate kinase Malate dehydrogenase Malic enzyme Carnitine acetyltransferase a The mean activity

* the standard

77.2 79.3 452.0 841.2 84.5 41.9

f + f i f f

9.0 9.6 70.1 93.0 9.8 4.4

76.3 78.6 655.0 875.1 52.0 40.8

zt zt i zt zt +

1.3 11.1 41.2 84.0 8.1 4.1

110.8 115.7 768.1 989.1 38.8 40.7

f + + i i zt

11.3 10.5 30.5 110.4 6.3 5.6

deviation.

preceding sexual maturity. Malic enzyme remained relatively constant in activity throughout the maturation period, whereas malate dehydrogenase was high in activity in the testis of newly eclosed males, declined in activity during the next few days of adult life, but increased greatly in activity late in the maturation period. Carnitine acetyltransferase increased in activity in the testis during the maturation period, but was less active in the mature testis than in the testis of 4day-old males. Enzyme activities in testis regions. Although the gametic content differences of sexually immature and mature testes were such as to make enzyme activities indicative of early and late stage gametes, respectively, enzyme assays on gamete preparations of greater purity were desirable to confirm the whole testis observations. Relatively pure gamete samples were acquired by dissection of the testes of IO-day-old males into three parts. The anterior region consisted of 70% spermatogonia and spermatocytes, and 35% early

spermatids; the middle region contained 100% mid and late stage spermatids; and the posterior region contained 100% mature sperm. Again, the ratio of spermatocytes to spermatogonia was at least 15: 1 (v/v). Glycolytic capacity, as indicated by hexokinase and pyruvate kinase activities, and Krebs cycle capacity, assessed by citrate synthase activity, were lowest in the early stage gametes of the anterior testis region and highest in the mature sperm of the posterior region (Table 7). Malate dehydrogenase activity, another Krebs cycle enzyme, appeared linked to amino acid metabolism. Both malate dehydrogenase and L-aspartate aminotransferase activities were high in spermatogonia and spermatocytes and in mature sperm. In contrast to L-aspartate aminotransferase activity, L-alanine aminotransferase activity was more than four times greater in mature sperm than in spermatogonia and spermatocytes, thus closely paralleling the activities of the glycolytic enzymes. The pattern for carnitine acetyltrans-

GEER et al.

Testis

and Thoracic

ferase (Table 7) was unique in that the enzyme was highest in activity in elongating spermatids, the middle region. However, the enzyme was also very active in mature sperm. There was no correlation between carnitine acetyltransferase activity and P-hydroxyacyl dehydrogenase activity. @Hydroxyacyl dehydrogenase, a fatty acid oxidation enzyme, was highest in activity in the anterior testis region and lowest in the posterior region. Malic enzyme, a NADP-dependent enzyme, was most active in the early stage gametes and least active in the late stage gametes (Table 7). The distribution of malic enzyme activity within the testis further suggested that the NADP-dependen\ enzymes are important to the growth and development of spermatocytes. Thoracic Tissue Metabolism Glycolytic and Krebs cycle enzymes. The glycolytic capacity of thoracic tissue from both immature and mature males appeared to be severalfold greater than that in the mature testis (Table 2). Phosphofructokinase and aldolase, two low activity enzymes in testicular glycolysis, were more than lo-fold more active in mature thoracic tissue than in the mature testis. Lactate dehydrogenase activity was extremely low in mature thoracic tissue

399

Tissue Maturation

and a-glycerophosphate dehydrogenase and pyruvate dehydrogenase complex activities were high (Table 2), suggesting that the Krebs cycle and cu-glycerophosphate dehydrogenase-oxidase cycle are the terminal steps of carbohydrate metabolism in D. hydei flight muscle. The glycolytic capacity of D. hydei male thoracic tissue increased 2- to 3-fold during adult maturation (Table 2), using phosphoglucomutase, hexokinase, and phosphofructokinase activities as indicators. These appeared to be the rate-limiting enzymes in the thoracic tissue glycolytic pathway. Similar increases in Krebs cycle and a-glycerophosphate dehydrogenase-oxidase cycle capacities, as reflected in the activities of a-glycerophosphate dehydrogenase and the Krebs cycle enzymes, were noted (Tables 2 and 3). By comparison, the Krebs cycle capacity of mature thoracic tissue appeared to be approximately three times that in the mature testis. In contrast, a-glycerophosphate dehydrogenase activity was almost loo-fold greater in mature thoracic tissue than in the mature testis. This reflects a major carbohydrate metabolism difference between the testis and thoracic tissues. NADP-dependent enzymes. The pentose shunt cycle appeared to be more important in the thoracic tissue of newly

TABLE 7 ENZYME ACTIVITIES IN Drosophila hydei TESTIS REGIONS Enzyme

Posterior

Hexokinase fructose glucose Pyruvate kinase Citrate synthase Malate dehydrogenase L-Alanine aminotransferase L-Aspartate aminotransferase Malic enzyme &Hydroxyacyl dehydrogenase Carnitine acetyltransferase o The mean activity

109.1 151.7 39.8 203.9 494.1 762.8 206.6 64.4 18.2 112.7

* the standard deviation.

* i + * + zt + f i +

15.7” 12.4 5.8 40.8 54.6 73.1 20.9 13.4 2.2 7.8

Middle

38.2 70.5 24.1 150.0 430.2 351.7 134.8 130.1 20.5 141.9

f 2.5 * 6.8 f 5.9 zt 26.3 f 63.4 zt 47.6 zi=13.8 l 19.0 + 1.2 * 7.1

Anterior

10.4 40.1 22.8 77.3 553.2 188.2 167.6 136.5 24.7 94.7

zt zt * zt + f zt zt + *

2.4 6.7 4.1 23.6 48.6 37.9 18.4 12.7 4.5 9.0

400

DEVELOPMENTAL BIOLOGY

eclosed males than in mature males (Table 4). Glucose-8phosphate dehydrogenase and 6-phosphogluconate dehydrogenase activities were moderately low in immature and mature thoracic tissues. Because the glycolytic capacity was much greater in mature thoracic tissue than in immature thoracic tissue, the importance of the pentose shunt cycle in carbohydrate metabolism appeared to diminish greatly during maturation. Malic enzyme and NADP-isocitrate dehydrogenase decreased markedly in thoracic tissue during maturation, perhaps reflecting a decline in synthetic capacity. In general, the NADPdependent enzymes were of less importance in mature thoracic tissue metabolism than in mature testis metabolism. Amino acid and fatty acid metabolism enzymes. Carnitine acetyltransferase was very active in the thoraces of both newly eclosed and mature males (Table 5), the activity being slightly smaller in mature thoracic tissues. ,&Hydroxyacyl dehydrogenase was moderate in activity in the mature male thorax, evidencing a minor role for fatty acid oxidation in mature thoracic tissue. The low levels of acetylCoA carboxylase in the thoraces of both test groups indicated that there was little fatty acid synthesis occurring at either age. Amino acid metabolism appeared prominent in immature and mature male thoracic tissues (Table 5). L-Aspartate aminotransferase was relatively active in immature male thoracic tissue, and increased slightly in activity during maturation, whereas L-alanine aminotransferase was very active in the thoracic tissues of newly eclosed males, but declined slightly in activity during maturation. By comparison acetyl-CoA carboxylase and L-aspartate aminotransferase activities were more prominent in the testis than in thoracic tissues; fi-hydroxyacyl dehydrogenase was somewhat more active

VOLUME 28, 1972

in thoracic tissue than in the testis; and carnitine acetyltransferase and L-alanine aminotransferase activities were very high in both testis and thoracic tissues. Carnitine acetyltransferase activity did not directly parallel the activities of the glycolytic enzymes during thoracic tissue maturation. The high activity of carnitine acetyltransferase in the immature thorax may have been related to the relatively high activities of L-alanine aminotransferase and P-hydroxyacyl dehydrogenase. This would not be surprising since carnitine acetyltransferase can facilitate fatty acid, amino acid, and carbohydrate metabolisms through the translocation of acetyl groups in mitochondria. Intermediate developmental stages. Examination of enzyme activities in thoracic tissue at intermediate stages of adult maturation (Table 6) revealed no unusual developmental patterns. The glycolytic enzymes, hexokinase and 3-phosphoglycerate kinase, reached near maximum levels by the midpoint of the adult maturation period, whereas the Krebs cycle enzyme, malate dehydrogenase, increased steadily in activity throughout the maturation period. In contrast, malic enzyme declined steadily in activity during the adult maturation period, and carnitine acetyltransferase activity remained relatively constant at all thoracic tissue maturation stages. The thoracic tissue hexokinase exhibited little, if any substrate specificity; e.g., essentially the same activities were observed when glucose and fructose were used as substrates. The testicular hexokinase activity, on the other hand, was greatest when glucose was employed as the substrate. Each enzyme activity that was examined had a testis maturation pattern that was different from the pattern of the same enzyme activity during thoracic tissue maturation (Table 6). This indicates that the enzyme activities in the two tissue

GEER et al.

types are subject to different factors during adult maturation.

Testis

and Z’horacic

regulating

DISCUSSION

The energy-yielding metabolism of Drosophila spermatozoa is similar to that of vertebrate spermatozoa. The present study and that by Blum et al. (1962) on honey bee sperm indicate that insect sperm, like vertebrate sperm (Mann, 1964), have the capacity to rapidly oxidize carbohydrate. Nevertheless, differences between mammalian and Drosophila spermatozoan metabolisms exist, and these have been incorporated into our model for metabolic changes during spermatogenesis. Drosophila spermatozoa apparently are unable to rapidly form lactic acid, a major anaerobic metabolic end product, for mammalian spermatozoa. Moreover, it is unlikely that Drosophila spermatozoa rely upon the a-glycerophosphate dehydrogenase-oxidase cycle as a means for aerobic NADH oxidation, a capacity that is well developed in Drosophila flight muscle. Although the a-glycerophosphate dehydrogenase-oxidase cycle appears operative to some extent, a-glycerophosphate dehydrogenase activity being low in both D. hydei and D. melanogaster spermatozoa (Geer, unpublished results), it appears not to be a major pathway in Drosophila spermatozoa. On the other hand, Blum and Taber (1965) reported higher a-glycerophosphate dehydrogenase activity for honey bee sperm, suggesting that the a-glycerophosphate dehydrogenase-oxidase cycle may be important to the metabolism of spermatozoa of insects other than Drosophila. In our model we suggested two possible anaerobic metabolic end products, alanine and acetylcarnitine. The high levels of Lalanine aminotransferase and L-aspartate aminotransferase observed for Drosophila, bovine (Flipse and Anderson, 1959; Flipse,

Tissue

Maturation

401

1960, 1962), and fish spermatozoa (Mounib, 1967) indicate that amino acid metabolism is important to a broad spectrum of spermatozoa. The distribution of Lalanine aminotransferase activity within the mature D. hydei testis parallels the activity distributions of the glycolytic enzymes, suggesting that this aminotransferase is functionally linked to glycolysis. Thus, Drosophila spermatozoa may anaerobically metabolize carbohydrate to pyruvate and then convert pyruvate to alanine. The alanine formed in this manner could later be metabolized aerobically by way of the Krebs cycle since the Krebs cycle is well developed in mature D. hydei gametes. In any event, Davey and Webster (1967) reported that a muco- or glycoprotein is secreted in the spermatheta of Rhodnius prolixus for use by stored sperm as an energy source, thus amino acid catabolism may be critical for maintenance of insect sperm while being stored in the female reproductive tract. Carnitine acetyltransferase, the enzyme that catalyzes the formation of acetylcarnitine from pyruvate, is extremely active in mature Drosophila (Geer and Newburgh, 1970) and rat sperm (Marquis and Fritz, 1965). Carnitine and/or its derivatives accumulate in the testes of Drosophila possessing functional mature spermatozoa but do not accumulate in the testes of sterile males (Geer and Newburgh, 1970). Consequently, acetylcarnitine may represent a storage form of “active” acetyl groups for Drosophila spermatozoa, a role that it has been suggested to play in flight muscle (Childress et al., 1966). Acetylcarnitine, like alanine, is readily metabolizable via the Krebs cycle. Drosophila spermatozoa may also differ from vertebrate spermatozoa in their capacity to oxidize lipid. Mammalian spermatozoa are capable of rapidly oxidizing phospholipid in the absence of carbohydrate (Mann, 1964) and Blum et al. (1967)

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DEVELOPMENTAL BIOLOGY

suggested this mode of energy metabolism for insect sperm after examining the lipid content of honey bee semen. The current study indicates that Drosophila spermatozoa have the capacity to oxidize fatty acids but suggests that this capacity is more limited than carbohydrate degradation, and is probably more limited than amino acid degradation. In oiuo observations of metabolite formation and turnover will be necessary to determine the relative importance of anaerobic and aerobic energy-yielding metabolism for Drosophila spermatozoa. In vitro enzyme activities suggest, as stated in our model, that Drosophila sperm are able to derive energy anaerobically by carbohydrate degradation and aerobically by either carbohydrate or amino acid degradation. The energy-yielding metabolism of early stage gametes differs markedly from that of late stage gametes in Drosophila. This is evidenced by changes in total activity for testicular enzymes at the time of sexual maturation plus the observation that a number of enzyme activities are differentially distributed in different regions of the testis. Glycolytic and Krebs cycle capacities are least in spermatogonia and spermatocytes but become increasingly greater until they reach their highest levels in mature sperm. Furthermore, fatty acid oxidation capacity, as indicated by /3-hydroxyacyl dehydrogenase activity, is greater in early stage gametes than in mature spermatozoa. The NADP-dependent enzymes appear to be important to the metabolism of developing gametes in another way. In Drosophila, the spermatocyte represents a period of extensive growth. Furthermore, Drosophila spermatids undergo dramatic structural alterations during maturation. Recent studies in this laboratory (Geer and Downing, 1972) have shown that lipid and protein are synthesized at rapid rates in the testis of D. hydei; consequently, high activity of the NADP-dependent

VOLUME 28. 1972

enzymes may be necessary to generate sufficient NADPH to support the synthetic processes associated with spermatogenesis. In any event, the prominence of the NADP-dependent enzymes in both immature and mature D. hydei testes reflects the fact that spermatogenesis is initiated before eclosion and continues during most of the life of the adult male. Distributions of the aminotransferases within the testis suggest that L-aspartate aminotransferase and L-alanine aminotransferase function in different ways during spermatogenesis. L-Alanine aminotransferase activity is much greater in mature spermatozoa than in early stage gametes. In contrast, L-aspartate aminotransferase activity is high in both early stage and mature spermatozoa and is least active in spermatids. L-Aspartate aminotransferase and malate dehydrogenase activities closely parallel each other during spermatogenesis, and their activities appear to be part of a metabolic pathway important to both early stage gametes and mature sperm. It is not possible at this time to state whether amino acid synthesis and/or degradation is the function of this pathway. Carnitine acetyltransferase activity seems most closely correlated to carbohydrate metabolism during spermatogenesis since it is more active in mature sperm than in spermatogonia and spermatocytes. It resembles the pyruvate metabolizing enzymes, pyruvate kinase, L-alanine aminotransferase, and the pyruvate dehydrogenase complex, in this respect. Nevertheless, maximum carnitine acetyltransferase activity is found in elongating spermatids, suggesting that the enzyme may play some metabolic role in spermatid differentiation. Carnitine acetyltransferase has been suggested to facilitate a variety of metabolic processes that utilize acetylCoA (Fritz, 1967), but there is no clue in the present study as to how the enzyme functions in the spermatid. Thus, as indicated in our model, the

GEER et al.

Testis

and Thoracic

activities of enzymes associated with metabolism, spermatozoan mature namely the glycolytic enzymes, Krebs cycle enzymes, and the aminotransferases, become prominent late in spermatogenesis. Carnitine acetyltransferase activity, as mentioned, becomes high in elongating D. hydei spermatids and remains high in mature sperm. On the other hand, the enzyme activities related to spermatocyte growth, e.g., the NADP-dependent enzymes, malate dehydrogenase, L-aspartate aminotransferase, and acetyl-CoA carboxylase, are high during the early stages of spermatogenesis. These changes in enzyme activity could be due to enzyme synthesis, activation, and/or inactivation. Little is known of the mechanism(s) controlling metabolic changes associated with spermatogenesis. It is known that development of the mature spermatozoan metabolism is dependent, at least indirectly, upon the action of Y-chromosome genes in Drosophila (Geer and Newburgh, 1970). Furthermore, since growth and differentiation of insect spermatogonia and spermatocytes is known to be hormone dependent (Takeuchi, 1969; Blaine and Dixon, 1970), it is likely that the metabolism of developing insect gametes is under hormonal control. Enzymatic changes that occur in D. hydei flight muscle during adult maturation reflect metabolic changes that are well documented in the Diptera and other insects (Biicher, 1965; Sacktor, 1965, 1970; Wyatt, 1968). In brief, during adult maturation in dipterans the number of thoracic mitochondria proliferates rapidly and mitochondria become much more highly structured (Lennie et al., 1967; Gregory et al., 1968; Walker and Birt, 1969). Thus, in mature Drosophila the flight muscle is densely packed with highly developed mitochondria and is well supplied with tracheoles (Williams and Williams, 1943). The levels of glycolytic and Krebs cycle enzymes, as shown in the current study, increase 2- to 3-fold from the time of

Tissue

Maturation

403

emergence to adult maturation and are severalfold higher than in the mature testis. There are some fundamental differences between the spermatozoan and thoracic tissue developmental patterns. One difference is that a highly functional a-glycerophosphate dehydrogenase-oxidase cycle develops during thoracic tissue maturation. In contrast, the cu-glycerophosphate dehydrogenase-oxidase cycle exists in Drosophila spermatozoa, but appears to be of minor importance. An equally important metabolic difference between testis and thoracic tissue development is the prominence of NADPdependent enzymes at all stages of testis maturation as contrasted to their declining activities in D. hydei thoracic tissue during maturation. The decline may reflect any of a number of possible metabolic changes. The NADP-dependent enzymes have been linked to lipid synthesis (Wise and Ball, 1964; Olson, 1966; Horie, 1968), and it is possible that a decline in activity of these enzymes represents a decline in thoracic tissue lipid synthesis. Nevertheless, acetyl-CoA carboxylase, the limiting enzyme in fatty acid synthesis, is low in activity in D. hydei thoracic tissue and recent radioisotope incorporation studies indicate that lipid is synthesized much more slowly in thoracic tissue than in the testis (Geer and Downing, 1972). Thus, the major functions of the NADP-dependent enzymes in flight muscle may not be in fatty acid synthesis. It seems more likely that malic enzyme and NADP-isocitrate dehydrogenase are involved in the cytoplasmic oxidation of substrate materials prior to the differentiation of thoracic mitochondria. The aminotransferases are very active at all stages of adult maturation of both testis and thoracic tissue. However, L-alanine aminotransferase is almost twice as active in the testis region containing mature spermatozoa than it is in thoracic tis-

404

DEVELOPMENTAL BIOLOGY

sue, and it is more than four times as active in mature spermatozoa than in the testis region possessing early stage gametes. This may reflect the proposed association of L-alanine aminotransferase activity with glycolysis in mature Drosophila spermatozoa and lack of such an association in thoracic tissue and early stage gametes. In conclusion, this study evidences energy-yielding metabolism differences between gametes at different stages of spermatogenesis. The metabolic pattern of the developing gametes is, in fact, distinct from the developmental metabolic pattern of thoracic tissue. We have proposed a model for gamete metabolism to relate our observations of testicular enzyme activities during sexual maturation to metabolic changes during spermatogenesis. It must be kept in mind that our model is based on in vitro enzyme activity determinations and that in vivo-in vitro activity differences may exist. We plan to test the model in future investigations. We are grateful to Dr. M. M. Green and Dr. James B. Boyd, Department of Genetics, University of California at Davis for their helpful criticisms during the preparation of this manuscript. We would also like to thank Dr. Green for supplying the Drosophila hydei Berendes wild-type strain. REFERENCES BAKER, W. K., and VON HALLE, E. S. (1953). The basis of the oxygen effect on x-irradiated Drosophila sperm. Proc. Nat. Acad. Sci. U.S.A. 39, 152-161. BEAMES, C. G., JR. (1963). Glycolytic enzymes of the aquatic snail, Physa hulei Lea. Comp. Biochem. Physiol. 8, 109-114. BEENAKKERS, A. M. T. (1969). Carbohydrate and fat as a fuel for insect flight. A comparative study. J. Insect Physiol. 15, 353-361. BEERMANN, W. (1965). Operative Gliederung der Chromosomen. Naturwissenschuften 52, 365-375. BEERMANN, W. (1967). Gene action at the level of the chromosome. In “Heritage from Mendel” (R. A. Brink, ed.), pp. 179-201. Univ. of Wisconsin Press, Madison, Wisconsin. BEIZENHERZ, G. (1955). Triose phosphate isomerase from calf muscle. Methods Enzymol. 1, 387-391. BERGMEYER, H. U., and BERNT, E. (1965a). Malic dehydrogenase. In “Methods of Enzymatic Analy-

VOLUME 28, 1972 sis” (H. U. Bergmeyer, ed.), pp. 757-760. Academic Press, New York. BERGMEYER, H. U., and BERNT, E. (1965b). Glutamate-oxaloacetate transaminase. In “Methods of Enzymatic Analysis” (H. U. Bergmeyer, ed.), pp. 837-842. Academic Press, New York. BLAINE, W. D., and DIXON, S. E. (1970). Hormonal control of spermatogenesis in the cockroach, Periplaneta americano (L). Can. J. Zool. 48, 283-287. BLUM, M. S., and TABER, S., III. (1965). Chemistry of the drone honey bee reproductive system. III. Dehydrogenases in washed spermatozoa. J. Insect. Physiol. 11, 1489-1501. BLUM, M. S., GLOWSKA, Z., and TABER, S., III. (1962). Chemistry of the drone honey bee reproductive system. II. Carbohydrates in the reproductive organs and semen. Ann. Entomol. Sot. Amer. 55, 135-139. BLUM, M. S., BUMGARNER, J. E., and TABER, S., III. (1967). Composition and possible significance of fatty acids in the lipid classes in honey bee semen. J. Insect Physiol. 13, 1301-1308. BRINK, N. D. (1968). Protein synthesis during spermatogenesis in Drosophila melanogaster. Mutation Res. 5, 1192-1194. BUCHER, T. (1965). Formation of the specific structural and enzymic pattern of the insect flight muscle. Zn “Aspects of Insect Biochemistry” (T. W. Goodwin, ed.), pp. 15-28. (Biochem. Sot. Symp. No. 25). Academic Press, New York. CHILDRESS, C. C., SACKTOR, B., and TRAYNOR, D. R. (1966). Function of carnitine in the fatty acid oxidase-deficient insect flight muscle. J. Biol. Chem. 242, 754-760. DAS, C. C., KAUFMANN, B. P., and GAY, H. (1964). Histone-protein transition in Drosophila melanogaster. I. Changes during spermatogenesis. Exp. Cell Res. 35, 507-514. DAVEY, K. G., and WEBSTER, G. F. (1967). The structure and secretion of the spermatheca of Rhodnius prolixus: A histochemical study. Can. J. Zool. 45, 653-657. DECKER, K. (1965). Acetoacetyl Coenzyme A. In “Methods of Enzymatic Analysis” (H. U. Bergmeyer, ed.), pp. 425-428. Academic Press, New York. DIPIETRO, D., and WEINHOUSE, S. (1960). Hepatic glucokinase in the fed, fasted and alloxan-diabetic rat. J. Biol. Chem. 235, 2542-2545. EPHRUSSI, B., and BEADLE, G. W. (1936). A technique of transplantation for Drosophila. Amer. Nat. 70, 218-225. FLIPSE, R. J. (1960). Metabolism of bovine semen. IX. Glutamic-oxaloacetic and glutamic-pyruvate transaminase activities. J. Dairy Sci. 43, 773-776. FLIPSE, R. J. (1962). Comments on certain aspects of the metabolism of spermatozoa. In “Spermatozoan Motility” (D. W. Bishop, ed.), pp. 133-136. Amer. Assoc. Adv. Sci. Publ. No. 72, Washington, D. C.

GEER et al.

Testis

and Thoracic

FLIPSE, R. J., and ANDERSON, W. R. (1959). Metabolism of bovine semen. XVII. Oxidative metabolism of glutamate. J. Dairy Sci. 52, 113-116. FRITZ, I. B. (1967). An hypothesis concerning the role of carnitine in the control of interrations between fatty acid and carbohydrate metabolism. Perspect. Biol. Med. 10, 643-677. GEER, B. W., and DOWNING, B. C. (1972). Changes in lipid and protein synthesis during spermatozoan development and thoracic tissue maturation in Drosophila hydei. Wilhelm Roux’ Arch. Entwicklungsmech. Organismen in press. GEER, B. W., and NEWBURGH, R. W. (1970). Carnitine acetyltransferase and spermatozoan development in Drosophila melanoguster. J. Biol. Chem. 245, 7179. GLOCK, G. E., and MCLEAN, P. (1953). Further studies on the properties and assay of glucose 6phosphate dehydrogenase and 6-phosphogluconate dehydrogenase of rat liver. Biochem. J. 55, 400408. GOEBELL, H., and KLINGENBERG, M. (1964). -DPNspezifishche Isocitrat-dehydrogenase der Mitochondrien I. Kinefishche Eigenschaften, Vorkommen und Funktion der DPN-spezifischen Isocitrat-dehydrogenase. Biochem. 2. 340, 441-464. GREGORY, D. W., LENNIE, R. W., and BIRT, L. M. (1968). An electron-microscopic study of flight muscle development on the blowfly Lucilia cuprina. J. Roy. Microsc. Sot. 88, 151-175. HENNIG, W. (1968). Ribonucleic acid synthesis of the Y-chromosome of Drosophila hydei. J. Mol. Biol. 38, 227-239. HESS, 0. (1966). Structural modifications of the Y chromosome in Drosophila hydei and their relations to gene activity. In “Chromosomes Today” (C. D. Darlington and K. R. Lewis, eds.), Vol. 1, pp. 167-173. Oliver and Boyd, Edinburgh. HESS, 0. (1967). Complementation of genetic activity in translocated fragments of the Y chromosome in Drosophila hydei. Genetics 56, 283-295. HESS, O., and MEYER, G. F. (1968). Genetic activities of the Y chromosome in Drosophila during spermatogenesis. Aduan. Genet. 14, 171-223. HORIE, Y. (1967). Dehydrogenase in carbohydrate metabolism in larvae of the silkworm, Bombyr mori L. J. Insect Physiol. 13, 1163-1175. HORIE, Y. (1968). The oxidation of NADPH by the soluble fraction of the fat body of the silkworm, Bombyr mori L. J. Insect Physiol. 14, 417-424. KORNBERG, A. (1955a). Lactic dehydrogenase of muscle. Methods Enzymol. 1, 441-443. KORNBERG, A. (1955b). Isocitric dehydrogenase of yeast. Methods Enzymol. 1, 699-714. LEFEVRE, G., JR., and JONSSON, U. B. (1964). X-ray induced mutability in male germ cells of Drosophila mekmogaster. Mutation Res. 1, 231-246. LENNIE, R. W., GREGORY, D. W., and BIRT, L. M. (1967). Changes in the nucleic acid content and

Tissue

Maturation

405

structure of thoracic mitochondria during development of the blowfly, Lucilia cuprina. J. Insect Physiol. 13, 1745-1756. LOWRY, 0. H., ROSEBROUGH,N. J., FARR, A. L., and RANDALL, R. J. (1951). Protein measurement with the Folin phenol reagent. J. Biol. Chem. 193, 265275. LOANING,K. G. (1961). Can Drosophila spermatozoa be used in studies of recovery processes? J. Cell. Comp. Physiol. 58, Suppl. 1, 197-201. MCCAMAN, R. E., MCCAMAN, M. W., and STAFFORD, M. L. (1966). Carnitine acetyltransferase in nervous tissue. J. Biol. Chem. 241, 930-934. MANN, T. (1964). “The Biochemistry of Semen and of the Male Reproductive Tract.” Wiley, New York. MARQUIS, N. R., and FRITZ, I. B. (1965). Effects of testosterone on the distribution of carnitine, acetylcarnitine, and carnitine acetyltransferase in the tissues of the reproductive system of the male rat. J. Biol. Chem. 240,2197-2200. MATSUHASHI, M., MATSUHASHI, S., and LYNEN, F. (1964). Zur Biosynthese der Fettsluren. V. Die Acetyl-CoA Carboxylase aus Rattenleber und ihre Aktivierung durch CitronensBure. Biochem. Z. 340, 263-289. MA~ENHEIMER, H. (1970). “Micromethods for the Clinical and Biochemical Laboratory,” pp. 137138. Ann Arbor Science Publ., Ann Arbor, Michigan. MOUNIB, M. S. (1967). Metabolism of pyruvate, acetate, and glyoxylate by fish sperm. Comp. Biothem. Physiol. 20, 987-992. NAJJAR, V. A. (lS48). The isolation and properties ot phosphoglucomutase. J. Biol. Chem. 175,281-290. OCHOA, S. (1955). “Malic” enzyme. Methods Enzymol. 1, 739-753. OLSON, J. A. (1966). Lipid metabolism. Annu. Reu. Biochem. 35,559-598. OSTER, I. I. (1961). On recovery in x-irradiated germ cells. J. Cell. Comp. Physiol. 58, Suppl. 1, 203207. SACKTOR, B. (1965). Energetics and respiratory metabolism of muscular contraction. In “Physiology of Insecta” (M. Rockstein, ed.), Vol. 2, pp. 483580. Academic Press, New York. SACKTOR, B. (1970). Regulation of intermediary metabolism, with special reference to the control mechanisms in insect flight muscle. Aduan. Insect Physiol. 7, 268-348. SHIOMI, T. (1967). Sensitivity differences in the successive stages of spermatogenesis in Drosophila after irradiation in nitrogen or air. Mutation Res. 4, 323-332. SHONK, C. E., and BOXER, G. E. (1964). Enzyme patterns in human tissues. I. Methods for the determination of glycolytic enzymes. Cancer Res. 24, 709-721. SOBELS, F. H. (1965). Radiosensitivity and repair in

406

DEVELOPMENTAL

BIOLOGY

different germ cell stages of Drosophila. In “Genetics Today” (S. J. Geerts, ed.), Vol. 2, pp. 227235. Pergamon Press, New York. SOBELS, F. H., and TATES, A. D. (1961). Recovery from premutational damage of x-irradiation in Drosophila spermatogenesis. J. Cell. Camp. Physiol. 58, Suppl. 1, 189-196. TAKEUCHI, S. (1969). Endocrinological studies on spermatogenesis in the silkworm, Bombyx mori L. Develop. Growth Differ. 11, 8-28. TIETZ, A., and OCHOA, S. (1958). Fluorokinase and pyruvic kinase. Arch. Biochem. Biophys. 78, 477493. TRAUT, H. (1964). Evidence for differential radiosensitivity rather than recovery in sperm samples from x-irradiated Drosophila melonogaster males. Genetics 50, 167-171. TROUT, W. E. (1964). Differential radiosensitivity as an explanation for so-called recovery in Drosophila sperm. Genetics 50,173-179. UNDERWOOD, A. H., and NEWSHOLME, E. A. (1965).

VOLUME

28.

1972

Properties of phosphofructokinase from rat liver and their relation to the control of glycolysis and gluconeogenesis. Biochem. J. 95, 868-875. WALKER, A. C., and BIRT, L. M. (1969). Development of respiratory activity and oxidative phosphorylation in flight muscle mitochondria of the blowfly, Lucilia cuprina. J. Insect Physiol. 15, 305-317. WARD, C. W., and SCHOFIELD, P. J. (1967). Glycolysis in Haemonchus contortus larvae and rat liver. Comp. Biochem. Physiol. 22, 33-52. WILLIAMS, C. M., and WILLIAMS, M. V. (1943). The flight muscles of Drosophila repleta. J. Morphol. 72, 589-599. WISE, E. M., JR., and BALL, E. G. (1964). Malic enzyme and lipogenesis. Proc. Nat. Acad. Sci. U.S.A. 52, 1255-1263. WYA~, G. R. (1968). Biochemistry of insect metamorphosis. In “Metamorphosis” (W. Etkin and L. I. Gilbert, eds.), pp. 143-184. Appleton-CenturyCrofts, New York.