Berndston 1977 44 818-33

William E. BerndstonMethods for Quantifying Mammalian Spermatogenesis: a Review

1977, 44:818-833.J ANIM SCI

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METHODS FOR Q U A N T I F Y I N G M A M M A L I A N SPERMATOGENESIS: A REVIEW 1

William E. Berndtson

Colorado State University 2, Fort Collins 80523

S U M M A R Y

Relatively large alterations in rates of spermatozoal production may be undetected un- less critical evaluation procedures are employed to assess spermatogenesis. Thus, a variety of techniques for quantifying spermatogenesis have been developed. These include the measurement of spermatozoal production in extirpated tissue by quantitative testicular histology or by homogenization of testicular tissue and subsequent enumeration of testicular spermatid reserves, measurement by determination of spermatozoal output in ejaculated semen or in the effluent obtained by cannulation of the male duct system of living subjects, and by several other specialized techniques. In this review, procedural details for each technique are described, and the limitations, advantages, disadvantages and basis for selection of each procedure is discussed. When used judiciously, these techniques constitute powerful tools for investigating the gametogenic activity of the testes. These techniques should be utilized whenever a critical assessment of the quantitative aspects of spermatogenesis is required. (Key Words: Spermatogenesis, Spermatozoal Production, Gametogenesis, Testis.)

I N T R O D U C T I O N

Definitive techniques are essential for the detection of all but extreme alterations in rates of spermatozoal production. To illustrate this point, the histological appearance of the testes of two rats is presented in figure 1. One of these rats was from a group of five untreated controls, while the other was from a similar group which received a high dose of exogenous

1 The author wishes to express his appreciation to Drs. R. P. Amann and G. lgboeli for their thoughtful comments and suggestions during the preparation of this manuscript.

2 Animal Reproduction Laboratory, Department of Physiology and Biophysics.

androgen. Although the tissue from these rats was grossly similar, and no differences in testis weight were observed (3.6 vs 3.3 g, respectively), the number of preleptotene spermatocytes, pachytene spermatocytes and step 7 spermafids in stage VII of the cycle of the seminiferous epithelium was reduced an average of 17 to 20% in the treated rats (Berndtson et al., 1974). In another study, Amann and Almquist (1961b) collected semen from bulls for 20 weeks following unilateral vasectomy. When only two ejaculates were taken per week, the numbers of spermatozoa per milliliter and per ejaculate were within the normal range for intact controls. Because even such large deficiencies in spermatozoal production can easily be undetected, a variety of techniques for quantifying mammalian spermatogenesis have been developed. The purpose of this review is to describe the techniques which are currently available and their limitations, advantages, disadvantages and basis for selection.

THE CYCLE OF THE SEMINIFEROUS EPITHELIUM

To fully understand and Utilize techniques for quantifying spermatogenesis one must pos- sess a fundamental knowledge of the cycle of the seminiferous epithelium. The cyclic nature of spermatogenesis results from the fact that at any given area within the seminiferous tubules, a well-defined series of events occur, which follow each other in a precise, orderly sequence. These events occur at extremely well-timed intervals with respect to one another, and give rise to a precise number of distinct cellular associations, each consisting of one or two generations of spermatogonia, spermatocytes and spermatids. The sequence of events occur- ring from the disappearance of a, given cellular association to its reappearance constitutes one cycle of the seminiferous epithelium (Clermont, 1972).

Fixed numbers of cellular associations arise 8 1 8 JOURNAL OF ANIMAL SCIENCE, Vol. 44, No. 5 (1977)



QUANTIFICATION OF MAMMALIAN SPERMATOGENESIS 819

Figure 1. Representative testicular tissue from untreated control animals (A) and rats treated with a large dose of exogenous androgen for 56 days (B).

within the germinal ep i the l ium because each germ cell, excep t certain spermatogonia , must divide at a precise interval fo l lowing its own fo rma t ion to yield specific daughter cells. The length of this interval varies for each type of

cell, bu t is relatively cons tan t within species. Figure 2 has been prepared to i l lustrate the con- cept of the cycle o f the semini ferous ep i the l ium and fo rma t ion of a specif ic n u m b e r of dis t inct cellular associations. This mode l is hypo the t i ca l and should no t be taken as depic t ing s tem cell renewal (de Rooij , 1968; de Rooi j and Kramer , 1968; Huckins, 1971a, b, c; C le rmont , 1972) or the sequence o f events during the in i t ia t ion of spermatogenesis at puber ty (C le rmon t and Perey, 1957b; Abde l -Raouf , 1960, 1961; At ta l and Couro t , 1963; Huckins, 1965; C o u r o t et al., 1970).

A n o t h e r feature responsible for dis t inct cellular associations is the synchronous division of cells of the same type in the same area of a semini ferous tubule in mos t mammals . The causes of this synchrony are n o t known, but intercel lular bridges jo ining groups of spermatogonia, spermatocy tes or spermat ids have been

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Figure 2. A series of hypothetical cellular divisions representing spermatogenesis is depicted along a time axis. For this model it is assumed that: 1) A cells divide 4 days after their own formation to produce type A and B cells, 2) B cells divide every 2 days to form C cells, 3) C cells divide i day after their own formation to yield D cells, 4) D cells divide after 1 day to produce E cells, 5) E cells divide after 2 days to form F cells which are released 4 days later as the final product of this process. Although 2 cells result from each division, only one mem- ber of each pair of identical daughter cells is shown. It can be seen that starting from a newly formed A cell, shown on day 1, 14 days are required to produce a mature F cell, i.e., the duration of spermatogenesis in this example is 14 days. Once the first generation of F cells is released, distinct cellular associations are observed which reoccur at 4-day intervals. Thus, one cycle of the seminiferous epithelium lasts 4 days, and 3.5 cycles (14 days) are required to complete spermatogenesis. Three distinct cellular associations can be distinguished. These consist of cells ABEF, ACF, and ADF, respectively. These associations have been utilized as criteria for dividing the cycle of the seminiferous epithelium into three stages, designated by Roman numerals I, II and III.



820 BERNDTSON

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observed in many mammals (Dym and Fawcet t , 1971; Huckins and Oakberg, 1971), and may be one means for assuring synchronous development (Fawcet t e t al., 1959; Huckins, 1971b; Huckins and Oakberg, 1971). This synchrony of differentiation also is responsible for the wave of the seminiferous epithelium, i.e., the distr ibution of cellular associations along the length of a seminiferous tubule at any given time (Curtis, 1918; Perey e t al. , 1961; Courot e t al. , 1970). The typical arrangement of cellular associations along the length of a seminiferous tubule of the rat is shown in figure 3. A wave of spermatogenesis has been identif ied in several other species, including the mouse, guinea pig, rabbit, bull, ram, boar, dog and cat (Von Ebner, 1871, 1888; Benda, 1887; Kramer, 1960; Hochereau, 1963). The physiological sig-

nificance of the wave is unknown, but its presence is helpful in that it reduces the presence of more than one cellular association when a seminiferous tubule is viewed in cross section, or when spermatogenesis is s tudied in whole mounts of seminiferous tubules (Clermont and Bustos-Obregon, 1968; Huckins and Kopriwa, 1969). A wave is not clearly established in man (Clermont, 1963 ; Steinberger and Tjioe, 1968). Rather, specific cellular associations appear in irregular zones (Heller and Clermont, 1964) as shown in figure 4, and cross sections of seminiferous tubules contain mixtures of cells in different stages. This fact has made quanti ta- t ion of spermatogenesis in man more difficult than in most other mammals (Steinberger and Tjioe, 1968). A similar arrangement exists in the cock (Clermont, 1958; Courot e t al. , 1970) and possibly in the stallion (Ellery, 1971).

Figure 3. The typical arrangement of cellular associations along the length of a seminiferous tubule of the rat is shown. Stages of the cycle of the seminiferous epithelium (designated by Roman numerals) are observed in a descending order distally from the attachment to the rete testis. Disruptions in this descending order (modulations) are depicted in the shaded area. A series of adjacent segments containing all stages of the cycle constitutes a wave of the seminiferous epithelium. The limit of one wave is denoted by arrows. (Adapted from Perey et al., 1961).

METHODS FOR D IST INGUISHING STAGES OF THE CYCLE OF THE

SEMINIFEROUS EPITHELIUM

The ability to divide the cycle of the seminiferous epithelium into identifiable compon- ent stages is an impor tan t prerequisite for most quantitative investigations of spermatogenesis. In figure 2, stages were designated solely upon the types of cells observed in a given association. If one employed no additional criteria, the number of stages would he constant within species. In actual practice, several o ther criteria are used, and the number of identifiable stages for a given species varies according to the criteria employed. Consequently, stages of the cycle of the seminiferous epithelium are arbi- trary, man-made divisions of a cont inuous pro-




Figure 4. The distribution of cellular associations (designated by Roman numerals) along the length of a seminiferous tubule of man is depicted. Although this distribution is hypothetical, the arrangement is representative of the facts. (Adapted from Heller and Cler- mont, 1964).

cess, rather than a biological constant for the species.

Two basic approaches have been used to identify stages of the cycle of the seminiferous epithelium. The first relies upon changes in the shape of the spermatid nucleus, t h e occurrence of meiotic divisions and the arrangement of spermatids within the germinal epithelium (Curtis, 1918; Roosen-Runge and Giesel, 1950; Ortavant, 1959). Because the specific arrangement of the germ cells, particularly elongated spermatids, is an important criterion, this will be designated the " tubular morphology" system. The second approach is based primarily upon changes in the acrosomic system and

morphology of developing spermatids (Leblond and Clermont, 1952), and will be designated the "acrosomic" system. Both the tubular morphology and acrosomic systems have been utilized to distinguish stages of the cycle of the seminiferous epithelium for many species, and a list of selected references which provide detail- ed criteria for distinguishing stages is sum- marized in table 1.

One advantage of the acrosomic system is greater ref inement due to the larger number of stages which are identified; although long stages identified by the tubular morphology system have occasionally been subdivided to overcome this objection (Hochereau-de Reviers, 1970). In addition, accurate identif icat ion of stages is often possible with the acrosomic system even after drastic reductions in the number of spermatids (Berndtson and Desjardins, 1974a). Elongated spermatids are often highly suscepti- ble to t reatment effects (Eschenbrenner e t al . ,

1948; Clermont and Morgentaler, 1955; Cler- mont and Harvey, 1967; Berndtson and Desjar- dins, 1974b; Berndtson e t al . , 1974). Thus, it is not uncommon to have a complete loss of elongated spermatids and marked reductions in the number of young spermatids following experimental t reatment of the testes. In such cases, the tubular morphology system cannot be used without adopting new criteria, which

TABLE 1. SUMMARY OF REFERENCES IN WHICH PROCEDURES FOR THE IDENTIFICATION OF STAGES OF THE CYCLE OF THE SEMINIFEROUS EPITHELIUM

OF DIFFERENT MAMMALS ARE DESCRIBED

Species System Reference

Boar Tubular morphology Ortavant (1959) Boar Tubular morphology Swierstra (1968a) Bull Tubular morphology Ortavant (1959) Bull Tubular morphology Amann ( 1962a) Bull Tubular morphology Swierstra (1966) Bull Tubular morphology Hoehere'au-de Reviers (1970) Bull Acrosomic Berndtson and Desjardins (1974a) Coyote Tubular morphology Kennelly (1972) Dog Tubular morphology Foote et al. (1972) Guinea pig Acrosomic Clermont (1960) Hamster Aerosomic Clermont (1954) Man Acrosomic Clermont (1963) Monkey Acrosomic Clermont and Leblond (1955, 1959) Mouse Acrosomic Oakberg (1956) Rabbit Tubular morphology Swierstra and Foote (1963) Ram Acrosomic Clermont and Leblond (1955) Ram Tubular morphology Ortavant (1959) Rat Acrosomic Leblond and Clermont ( 1952) Rat Acrosomic Clermont and Percy ( 1957a) Stallion Tubular morphology Swierstra et al. (1974)



822 BERNDTSON

generally cause pooling of several stages and hence, reduced precision. However, if young spermatids are present, even in relatively small numbers, the acrosomic system still can be applied. If these cells are lost, one must also develop new criteria, but this would require a much more severe insult to the testes than that required to preclude use of the tubular morphology system. Thus, the acrosomic system offers greater refinement, and the opportunity for application under numerous conditions during which the tubular morphology system would not be usable. Disadvantages include the fact that because one must examine spermatid cytology, rather than tubular histology, identification of at least certain of the stages is more difficult and time-consuming. Furthermore, since fixation and staining procedures alter the appearance of the germ cells (Clermont, 1972), it may be necessary to utilize the same procedures if existing descriptions (table 1) are to be employed. Finally, as the number of identifiable stages is increased, the frequency of a given stage is likely to be reduced. Thus, more time may be required to locate the desired number of tubules in any given stage.

PROCEDURES FOR QUANTIFYING SPERMATOGENESIS

Quantitative techniques are most often used to measure relative changes in spermatozoal production rates, absolute spermatozoal production or to investigate precise spermatogenic events. Procedures for each of these applica- tions follow.

Relative Changes in Spermatozoal Production Rates

Establisbment of Relative Spermatozoal Pro- duction Rates from Testicular Spermatid Re- serves. Relative changes in spermatozoal production rates can be established by direct comparison of testicular spermatid reserves, i.e., the number of spermatids per testis, per pair of testes, or per Unit weight of testicular parenchyma. The technique for determining testicular spermatid reserves involves homogenization of testicular tissue and subsequent enumeration of elongated spermatids by hemocytometry (Amann and Almquist, 1961a, 1962; Orgebin- Crist, 1968; Amann and Lambiase, 1969). Be- cause the time required for these cells to be transformed into spermatozoa is relatively constant for members of the same species, relative rates of spermatozoal production can be deter-

mined by comparing the size of the spermatid reserves in control and experimental animals. The technique is relatively simple and less time- consuming than histological procedures, but practice is required to distinguish elongated spermatid nuclei from tissue debris (Amama and Almquist, 1961a; Amann and Lambiase, 1969). The technique can be utilized for comparing spermatozoal production rates among members of the same species in the absence of procedures for identification of stages of the cycle. Comparisons among species are not valid, because the number of days' spermatozoal production represented by the elongated spermatids in testicular homogenates may differ.

Determination of Relative Spermatozoal Production Rates by Testicular ttistology. Cler- mont and Morgentaler (1955) proposed a method for measuring relative spermatozoal production rates in which one stage of the cycle was chosen as representing spermatogenesis as a whole (Clermont and Harvey, 1967). Germ cell and Sertoli cell nuclei or their identifiable frag- ments were then enumerated in round, seminiferous tubular cross sections of both experimental and control animals. The number of nuclei counted was designated the crude count, which required adjustment for two factors. The first correction was based upon the fact that nuclei of larger diameter are more likely to be counted in sections of any given thickness. Thus, although two different cell types may be present in equal numbers, the crude cell counts will differ if their nuclei are of different size.

This is exemplified in the data of table 2, which contains typical crude cell counts for germ cells in stage VII of the seminiferous epithelial cell cycle of the rat. From the crude counts it would appear that the testis contained more pachytene than preleptotene spermatocytes. This is impossible in the normal testis since pachytene primary spermatocytes evolve from preleptotene primary spermatocytes without cellular division. The problem resides in the fact that the pachytene primary spermatocyte nuclei are larger in diameter (table 2). Abercrom- bie (1946) developed a procedure by which crude counts can be adjusted to permit their direct comparison. This correction is valid for any spherical structure in a histological section. Abercrombie's formula is as follows:

True cell count = crude cell count • [(Section thickness (U)] /[Section thickness (~t) +

nuclear diameter (/~)]



QUANTIFICATION OF MAMMALIAN SPERMATOGENESIS

TABLE 2. MEAN SIZE AND NUMBER OF GERM CELLS PER STAGE VII SEMINIFEROUS TUBULAR CROSS SECTION OF THE RATa

823

Item

Cell type

Type A Preleptotene Pachytene Step 7 spermatogonia spermatocytes spermatocytes spermatids

Crude count 2.85 62.51 79.17 265.37 Nuclear dia. (/J) 6.8 6.01 8.29 6.94 Abercrombies' correction factor .37 .40 .325 .366 True count 1.05 25.0 25.73 97.13

aMean based upon 20 tubular cross sections from each of five rats. Sections were cut at 4/a thickness.

As anticipated, when the crude counts in table 2 were corrected by this procedure, the true counts for both the preleptotene and pachytene primary spermatocytes were similar (table 2).

To correct crude cell counts the thickness of the histological sections must be known. Gen- erally, this has been assumed to equal the microtome setting, but may be quite different and can vary among sections prepared in a series at a given setting. Since variation in section thickness can introduce errors in the cell correction factors, it has been recommended that the thickness of each section be determined by interference microscopy (HalMn, 1962; Amann, 1970a).

Crude cell counts must also be corrected for changes in tubular dimensions as a result of experimental treatment or histological procedures. Let us assume that an experimental treatment produced a 50% reduction b o t h in the spermatozoal production rate and the length of the seminiferous tubules, while tubular diameter was unchanged. The number of germ cells per tubular cross section should be identical to that of control animals, and a comparison of the mean number of germ cells per tubular cross section would reveal no treatment effect. This problem can be prevented by use of a Sertoli cell correction factor (Clermont and Morgentaler, 1955; Lino, 1971). This correction is based on the finding that Sertoli cells rarely divide in adult animals and are extremely resistant to most factors which adversely affect the germ cells. Thus, these cells serve as a point of reference for changes in germ cell numbers. The number of germ cells is adjusted to a per- Sertoli-cell basis. Frequently the number of Sertoli cells per tubular cross section in control animals serves as a base to which other cell counts are related. Assumptions inherent in the use of a Sertoli cell correction factor are: 1) the

number of Sertoli cells is relatively constant for normal animals of the same age and species, and 2) Sertoli cells are not destroyed by the experimental treatment.

The number of Sertoli nuclei should be corrected for nuclear diameter before use as a correction factor. However, the shape of Sertoli nuclei is irregular and varies at different stages of the seminiferous epithelial cell cycle (Elft- man, 1950; Leblond and Clermont, 1952). Since Abercrombie's formula is for use with spherical structures, correction of Sertoli cell counts is difficult. Lino (1971) developed a formula for calculating Sertoli cell nuclear vol- umes in the ram. However, a simpler solution is to count only those Sertoli cell nuclei which contain a nucleolus (Clermont and Harvey, 1967; Berndtson e t al., 1974). The latter are spherical, thus enabling adjustment by Aber- crombie's formula. This approach requires the assumption that the proportion of Sertoli cells with stainable nucleoli is not affected by experimental treatment.

The choice between histological or homogenization techniques for quantifying relative changes in spermatozoal production depends largely upon the nature and anticipated severity of the effects. Testicular spermatid reserves can be determined quickly, the procedures are much simpler and less tedious than histological methods, and a larger sample of the testis can be analyzed. Secondly, since elongated spermatids are the last cells to be formed, alterations in spermatogenesis at any point should ulti- mately be reflected in a change in the spermatid reserves. One limitation of these techniques is that they would not be useful in establishing the relative severity of treatment effects on ear- lier spermatogenic events or for monitoring the recovery of spermatogenesis after a severe insult to the testis in the absence of elongated sperma-



824 BERNDTSON

tids. For example, if only spermatogonia remained following a severe insult to the testis, it would be several weeks before elongated spermatids reappeared. However, histological changes would provide evidence of recovery, or the lack of it, throughout this period.

One disadvantage of histological procedures is that elongated spermatids cannot be counted accurately, because of their irregular shape and their arrangement within the germinal epithelium, and procedures for converting crude counts to true counts have not been developed. Thus, these techniques are only useful for quantifying spermatogenesis up to the elongation of spermatids. Cellular losses during subsequent steps of spermiogenesis would not be detected. These techniques are also tedious and time consuming, but provide some insight as to which classes of germ ceils (i.e., spermatogonia, spermatocytes, spermatids) are affected by a given treatment. They are also applic- able when elongated spermatids are absent, especially if the acrosomic system for determining stages of the cycle of the seminiferous epithelium is used.

The nature of the testicular response following experimental treatment can also influence the choice of quantitative technique for assessing spermatozoal production. With histological procedures, cross sections of seminiferous tubules are considered as representative of the testis as a whole. This is valid provided all seminiferous tubules are affected uniformly and the histological sections are separated by a reason- able distance. The seminiferous tubules of most mammals are highly convoluted (Curtis, 1918; Clermont and Huckins, 1961;Swierstra, 1968a; Amann, 1970a). Thus, cross sections of the same tubule may appear frequently in a section of testicular tissue from a single location. Therefore, Amann (1970a) has recommended that "at least five tissue sections separated from each other by a minimum distance of 1,000/a" be utilized for histological evaluations. This recommendation seems especially valid since all tubules need not be affected uniformly. Allan- son e t al. (1935) and Cutuly (1941) indicated that hypophysectomy of male guinea pigs was followed by the complete arrest of spermatogenesis in some seminiferous tubules, while spermatogenesis appeared grossly normal in others. Under such circumstances it would be exceedingly difficult to obtain representative sampling for histological procedures. A similar problem would exist any time a given treat-

ment, such as removal of a biopsy specimen, produced a localized response. Under such circumstances homogenization of the entire testis and subsequent enumeration of testicular spermatids would appear superior. Nonethe- less, comparison of testicular tissue removed from several locations within the same testis of normal animals have generally shown similar histological results (Amann, 1962a; Kennelly and Foote, 1964). Thus, if individual histological sections are separated by about 1,000 /a, there is generally no need to sample more than one location within a given testis.

Swierstra e t al. (1964) reported that intra- venous injection of Fungizone in male rabbits caused a delay in the release of spermatids. One must expect that if the rate of spermatozoal production was not affected, spermatid reserves would increase simply due to delay in spermatozoal release and might be interpreted as a true increase in spermatozoal production. This situa- tion is probably uncommon, but could only be detected by histological methods.

Neither the histological nor homogenization technique for assessing relative rates of spermatozoal production is clearly superior in all instances. Moreover, it is often impossible to predict the nature or the severity of a given treatment in advance. Under such circumstances it is advisable to process testicular tissue so that either or both procedures may be utilized. Fur- thermore, the technique to be utilized for quantifying spermatogenesis should be considered during the design of an experiment, since method of evaluation will, in part, determine the time(s) at which tissues should be taken for evaluation. To illustrate this point, let us assume we wish to utilize the size of testicular spermatid reserves to determine if the single injection of a specific drug has an influence on spermatozoal production. It must be recognized that several weeks are required for spermatozoa to form from spermatogonia (Clermont, 1972). Thus, if the drug exerts specific effects on spermatogonia, it will be several weeks before the size of the spermatid reserves is affected. In contrast, if the drug influenced only elongated spermatids, changes in spermtid reserves might be evident almost immediately, and normal numbers of these cells could be restored within a few days. Obviously, tissues must be taken at suitable intervals after treatment to permit detection of effects at any point in spermatogenesis by the technique selected. By using both the histological and spermatid reserve




methods, more cell types could be evaluated in tissues obtained at any given time. This would allow one to maximize the information gained from a single animal, thereby reducing the cost and labor to resolve this type of question.

Quantification of Relative Changes in Rates of Spermatozoal Production in Species in which a Wave of Spermatogenesis is Absent. Because a wave of spermatogenesis is absent in the cock (Clermont, 1958; de Reviers, 1968; Courot et al., 1970), man (Clermont, 1963) and possibly in the stallion (Ellery, 1971), cross sections of seminiferous tubules generally contain more than one cellular association. Thus, application of the afore- mentioned histological procedure for quantifying changes in relative rates of spermatozoal production is difficult. Steinberger and Tjioe (1968) have proposed an alternative procedure for use in evaluating" testicular tissue from the human. With this technique, a histological section of an entire biopsy specimen was projected on a sheet of paper and the outline of each tubule traced, numbered, and the circumfer- ence measured. Tubules were identified with a microscope. Those cut at an oblique angle showing no lumen were disregarded. The various germ cells then were identified (Clermont, 1963) and counted. Counts were expressed per unit length (33.3 #) of the seminiferous tubules. A random sampling of 25 tubules provided a reliable estimate for the entire biopsy specimen. Cell counts were similar for three normal individuals, while a significant reduction in the number of specific germ ceils was detected in oligospermic males. Thus, this technique would appear to permit the quantification of relative changes in spermatogenesis, without the need for distinguishing stages of the cycle, although identification of cell types is required. The technique can be performed on a biopsy specimen, and can provide specific insight as to the types of cells which may be affected under pathological conditions. How- ever, the reliability of estimates from only 25 tubules in a single biopsy specimen was based upon comparisons among only three normal individuals. Thus, sampling error and error due to shrinkage of the tissue due to histological processing must be considered as potential sources of error. This technique has not been evaluated for use in other species. Nonetheless, it may be useful in instances in which a histological method is preferred, although a distinct wave of spermatogenesis is lacking.

Determination of Absolute Spermatozoal Production Rates

Calculation of Daily Spermatozoal Produc- tion from Testicular Spermatid Reserves. If one knew the size of the spermatid reserves and how many days were required for these spermatids to form spermatozoa, daily spermatozoal production could be readily calculated. For example, if the testes contained 50 billion elongated spermatids, and 10 days were required for these to be released as spermatozoa, daily spermatozoal production would equal 5 billion. The critical point is to establish how many days' spermatozoal production are represented by the testicular spermatid reserves. This interval has been designated the "time divisor". Initially the time divisor was assumed to equal the sum of the duration of the stages of the cycle of the seminiferous epithelium in which elongated spermatids were present. This required the assumption that elongated spermatids were resistant to damage during homogenization. Orgebin-Crist (1968) concluded that elongated spermatids in testicular homogenates of the rabbit represented spermatids from stage V on, in the classification scheme of Swierstra and Foote (1963). However, elongation is com- pleted in stage II. Thus, elongated spermatids in late stage II and stages III and IV must have been damaged by homogenization. In support of this, Amann and Lambiase (1969) also presented evidence that elongated spermatids in stage IV do not survive homogenization. Be- cause of this, they determined the time divisor in the rabbit from: 1) the sum of the duration of those stages in which elongated spermatids were observed histologically, 2) the interval from the first appearance of labeled spermatids in testieular homogenates to the time these cells were first released from the seminiferous tubules, and 3) the interval between the appearance of the first spermatids in testicular homogenates up to the time labeled spermatids first entered the caput epididymidis. The time divisors averaged 3.5, 3.0 and 3.5 days, respectively. These investigators suggested that a pooled value of these estimates might give the best available time divisors and concluded that: "The accuracy of daily sperm production values calculated from testicular spermatid reserves is dependent upon: (a) the accuracy of the time divisor, (b) the efficiency of homogenization,

(c) the extent of destruction of elongated spermatids during homogenization and (d) the



826 BERNDTSON

variability associated with hemacytometric counting." Accuracy also depends upon the rate of cellular degeneration during the late steps of spermiogenesis. However, daily spermatozoal production can be determined much easier from testicular spermatids than by histological procedures, particularly when used for those species for which the time divisor has been established.

Several assumptions are required for determining spermatozoal production rates from testicular spermatid reserves. The first is that all elongated spermatids will form spermatozoa. This assumption is difficult to test, and must be considered as a potential source of overestima- tion, since degeneration of germ cells is recognized as a "normal" feature of spermatogenesis in many species (Amann, 1962b; Barr et al., 1971 ; Huckins, 1972; Roosen-Runge, 1973). A second assumption is that the length of one cycle of the seminiferous epithelium is constant. Dif- ferences in the duration of one cycle have been found among pubertal and post-pubertal animals (Huckins, 1965), animals of different strains (Clermont, 1972), and occasionally among members of the same strain (Amann and Lambiase, 1969). However, differences in the duration of one cycle of the seminiferous epithelium among members of the same species have generally been small, and no factors have been found to alter the length of the cycle.

Determination of Daily Spermatozoal Pro- duction from the Volumetric Proportions of the Testicular Elements. To estimate daily spermatozoal production from the volumetric proportions of the testicular elements, testes are removed and weighed. This weight must be corrected for the weight of the tunica albuginea and, if present, the mediastinum. Appropriate samples of parenchyma then are removed and weighed in air and in water. The density of the tissue is determined as mass divided by volume, with volume determined by Archimedes principle (i.e., the difference in the weight of an object in air and in a liquid equals the weight of the fluid displaced). Volume and density are again determined after histological processing and paraffin impregnation by weighing in air and in absolute alcohol (Swierstra, 1966, 1968b). The amount of shrinkage due to histological processing, which is directly proportion- al to the increase in density of the tissue, is then determined. Volumetric proportions of the testicular elements are determined by Chalkley's method (Chalkley, 19r Eschen-

brenner et al., 1948). With this procedure, pointers are attached to the ocular of a microscope, fields are selected at random, the structures under the pointers brought into focus and recorded. This process is repeated for a large number of fields. The frequency with which a

given structure appears under the pointer is directly related to its relative volume in the testis. From the total volume of testicular parenchyma and the percentage of the parenchyma occupied by cells of a given type, the total volume of cells of a given type can be calculated. The number of these cells can in turn be calculated by dividing the total volume of these cells by the mean volume of a single cell. Since cell volume is measured after histological processing, appropriate correction must be made for shrinkage. The underlying assumption is that shrinkage is uniform for all elements of the same testis. Shrinkage must be determined for each individual testis. Because the shape of elongated spermatids is irregular and their volume difficult to calculate, round spermatids or primary spermatocytes are usually employed. If primary spermatocytes are used, consideration must be given to the fact that each should divide to produce two secon- dary spermatocytes, which in turn produce four spermatids. Foote (1962) provided the following formula for estimating daily spermatozoal production from the number of primary spermatocytes per testis.

Estimated daily spermatozoal production per testis = [(number of spermatocytes per testis)

(% of spermatocytes completing spermatogenesis) (four spermatozoa from one primary spermatocyte)l /

(number of days required for evolution of primary spermatocytes)

Swierstra (1966) utilized the volumetric proportions of all round spermatids or round spermatids in "stage I" to determine spermatozoal production rates by the bovine testis. In principle, spermatozoal production rates can be determined from any given cell type. However, as less differentiated cells are utilized, one must take into account the number of divisions which take place to form spermatids, and that spermatozoal production may be increasingly overestimated, since one cannot account for subsequent cellular degeneration.

Measurement of Daily Spermatozoal Produc- tion from the Number of Spermatids in Histo- logical Sections. Amann and Almquist (1962)




proposed a technllque for calculating daily spermatozoal production from the number of round spermatid nuclei in histological sections. Round spermatids were enumerated in cross sections of seminiferous tubules in stage I according to the tubular morphology system, the percentage of the testis occupied by seminiferous tubules was determined by Chalkley's procedure (Chalkley, 1943) and the mean area of stage I tubular cross sections was determined. Daily spermatozoal production then was calculated by the following equations:

Corrected testis volume = [(testis weight)(shrinkage correction) (100 - %

tunica albuginea and mediastinum)]/(testis density)

Daily spermatozoal production = [(corrected testis volume) (% seminiferous

tubules in testis) (corrected no. of spermatids/stage I tubular cross section)] /[(duration of one cycle

of seminiferous epithelium) (area of stage I tubular cross section) (section thickness)]

As with all histological procedures for determining spermatozoal production rates, this method is exceedingly tedious and time-consuming, and is subject to potential error from the assumption that shrinkage due to histological processing is similar for all testicular elements, and because losses due to degeneration at later phases of spermatogenesis would not be detected. Because adverse treatments frequently cause greater degeneration among the mature germinal cells, this technique would be less desirable than the homogenization procedure when an adverse testicular response is anticipated. All procedures should be equally valid for estimating spermatozoal product ion of normal animals.

Daily Spermatozoal Output. One.of the old- est and simplest approaches for quantifying spermatozoal production is by measuring spermatozoal output . Spermatozoal output re- presents the number of spermatozoa which can actually be harvested from an animal, in contrast to spermatozoal production, which is the number of spermatozoa actually produced. Spermatozoal output has generally been determined from the number of spermatozoa re- covered in ejaculated semen. The major advantage of this approach for estimating spermatozoal product ion is that the reprodUctive capacity of an animal is not impaired. Thus, the information can be used to select animals for breeding purposes and for studying changes with time in the same animal. However, ade-

quate experimental design is essential if spermatozoal ou tpu t is to provide a reliable estimate of spermatozoal product ion.

The influence of extragonadal sperm reserves must be considered in the design of an exper iment in which spermatozoal ou tput is to be used to estimate spermatozoal production. Extragonadal spermatozoa are spermatozoa within the male reproductive system but not within the testes. If an animal is sexually rested, the extragonadal sperm reserves will attain a maximum size. Subsequently, the number of spermatozoa which enter the reserves from the testis will be offset by equal losses of spermatozoa by resorption, micturi t ion or masturbat ion (Lino et al., 1967; Holtz and Foote , 1972; Amann et al., 1974; Amann and Lambiase, 1974). Because of such losses, spermatozoal output in ejaculated semen will const i tute an inadequate estimate of spermatozoal production if semen is collected infrequently. Thus, the interval between seminal collections must be sufficiently short to minimize spermatozoal losses of this type. Amann (1970a) suggested that "A uniform interval of 1 or 2 days, probably rarely more than 3 days, between semen collections might be considered for use with most species."

Spermatozoal output will also differ from spermatozoal product ion if changes in the size of the extragonadal sperm reserves over an experimental period are not taken into account . For example, the extragonadal sperm reserves of sexually rested, adult livestock animals are approximately as follows: dairy bulls, 70 billion (Almquist and Amann, 1961 ; Amann and Alm- quist, 1961a); stallions, 58 billion (Gebauer et al., 1974b), boars (epididymal reserves 72 hr after depletion), 166 billion (Swierstra, 1971); and rams (epididymal reserves), 160 billion (Ortavant, 1958). These values represent approximately one to two weeks' spermatozoal production. The extragonadal sperm reserves are reduced by ejaculation. In the bull, for example, approximate ly 50% of the extragonadal sperm can be removed by deplet ion (Alm- quist and Hale, 1956; Almquist and Amann, 1961). Thus, when sexually rested animals are placed on an intensive seminal collection schedule, spermatozoal ou tput will reflect both the rate of spermatozoal product ion and the extent to which the extragonadal reserves have been depleted. Thus, Ortavant (1952, 1959) proposed the following formula for determining average daily spermatozoal product ion:



828 BERNDTSON

X = m - [(QI - Qa) /n l , i n which

m = average quantity of spermatozoa collected daily during n days;

QI = sperm reserves in the tail of the epididymis at the beginning of the experiment;

0.2 = sperm reserves in the tail of the epididymis at the end of the experiment.

This equation is valid only when the interval between seminal collections is sufficiently short to minimize spermatozoal losses through resorption, micturition or masturbation. Apparently Ortavant considered that most spermatozoa come from the tail of the epididymis during ejaculation. However, because the number of spermatozoa in the vas deferens and ampulla may also change, it would be more accurate to substitute total extragonadal reserves for cauda epididymal reserves in this equation.

The most difficult problem in satisfying Ortavant's equation is the determination of epididymal or extragonadal sperm reserves, particularly at the beginning of an experiment. However, the correction for a change in the extragonadal sperm reserves will become in- significant if the collection period is very long or the change in the extragonadal reserves is small (Almquist and Hale, 1956; Ortavant, 1959). Thus, one of two approaches may be used: 1) collect semen for extended periods or 2) stabilize the extragonadal sperm reserves by collecting semen at the experimental frequency prior to the experiment. The first approach will generally be less advantageous because short- term responses to experimental treatment cannot be readily investigated. Furthermore, this approach suffers from increased costs, time and labor requirements, the possibility of confound- ing by seasonal changes in spermatozoal production in some animals and because of diffi- culty in maintaining adequate libido in all animals. Therefore, it has been suggested that extragonadal sperm reserves be stabilized by collecting semen at the experimental frequency for 7 to 10 days before collecting spermatozoal output data (Amann, 1970a). Although such an approach will generally be most advantageous, the possibility of a change in the size of the reserves due to experimental treatment cannot be excluded.

Proper procedures for the collection of ejaculated semen are also essential if spermatozoal output data are to be meaningful. The response to sexual preparation procedures varies greatly

among species. For example, the use of false- mounting or active restraint results in marked increases in the spermatozoal output of bulls (Hale and Almquist, 1960; Hafs et al., 1962; Almquist, 1973). In contrast, spermatozoal out- out is not influenced by such procedures in boars (Signoret, 1 9 7 0 ) o r stallions (Pickett and Voss, 1973). In the latter, sexual stimulation increases seminal volume without altering spermatozoal output. The increased volume is largely due to increased secretion of gelatinous material, which increases spermatozoal losses. Thus, individuals should learn and utilize opti- mal seminal collection techniques for the species under study. In addition, spermatozoal output data must be corrected for spermatozoal losses in the seminal collection equipment (Foote and Heath, 1963 ; Pickett et al., 1974).

From studies with the bull, it would appear that spermatozoal production rates are not influenced by the frequency of ejaculation (Amann, 1962a). Correlations between estimated daily spermatozoal production and daily spermatozoal output of different animals have varied, and include .87 (Amann and Almquist, 1962) and .79 (Amann et aL, 1974) for bulls on a frequent seminal collection schedule, while

Swierstra (1966) found no significant correla- tion for bulls electroejaculated every other day. Corresponding correlations for other species include: stallions .80 (Gebauer et al., 1974a); boars, .54 (Swierstra, 1968b); and rabbits, .75 (Amann, 1970b) and .86 (Lambiase and Amann, 1969). In general, it would appear that spermatozoal output may be utilized as a method for estimating spermatozoal production of animals from which spermatozoa can be collected if appropriate procedures are followed. How- ever, spermatozoal output provides only limited insight as to which spermatogenic events may be affected by a given treatment. If spermatozoal output is determined by collection of semen with an artificial vagina, all animals must have high libido, and procedures or treatments which may alter libido are precluded.

Cannulation o f the Male Duct Sys tem. Spermatozoal production rates have been deter-- mined by cannulating the fete testis (Amann et al., 1963; Voglmayr and Mattner, 1968; Vogl- mayr et al., 1972). In principle, this is ideal since all spermatozoa leaving the testis should be accounted for. This procedure is currently restricted to use in large animals. Voglmayr et al. (1966, 1967) reported that catheters in the ram remained patent for up to 20 days. A1-




though catheters in the rete testis of the bull remained patent for up to 43 days, spermatozoal output declined markedly after 7 to 12 days (Voglmayr et al., 1972). These and other data (Amann et al., 1974) support the conten- tion that the procedure may alter spermatogenesis, and that cannulation of the rete testis should only be considered for short-term studies. The procedure is also precluded in instances in which it is desirable not to impair the reproductive life of the individual.

Cannulae in the vas deferens appear to produce fewer surgical complications and remain patent longer than those in the rete testis (Bennett and Rowson, 1963;Wierzbowski and Wierzchos, 1969; Setchell et al., 1971; Sexton et al., 1971; Holtz and Foote, 1974;Deutscher et al., 1974), and can be utilized for livestock species and the larger laboratory animals (Holtz and Foote, 1974). The major disadvantages are that: 1) spermatozoal resorption may occur in the epididymis, 2) the procedure is precluded in valuable animals in which permanent altera- tion of reproductive capacity is undesirable, and 3) one gains only limited insight as to which spermatogenic events may be influenced by a given treatment.

Anastomosis of the Vas Deferens to the Urinary Bladder. Vreeburg et al. (1974) recent- ly described a method for anastomosing the vas deferens to the urinary bladder in the rat. Urine then was collected and spermatozoa enumerated. Of 10 rats utilized, one failed to emit spermatozoa in the urine. However, a constant number of spermatozoa were voided in the urine of other rats for over 4 months. Thus, this procedure could be utilized for monitoring changes in spermatozoal production over time. This technique shares many of the disadvantages common to spermatozoal output or cannulation studies. First, one must assume that the procedure does not impair spermatozoal production. Also, spermatozoal output is subject to error from spermatozoal resorption, changes in the size of the extragonadal sperm reserves and the efficiency of spermatozoal recovery in the urine. Holtz and Foote (1972) could recover only about two-thirds of all rabbit spermatozoa sprayed into a cage designed to collect urine, and only one-half of these (one-third of the total) were detectable after processing with urine. Such low recovery clearly reduces the sensitivity of such procedures for studying spermatozoal production. Thus, this technique will require further evaluation before

its use can be recommended. Quantitation of Changes in Spermatozoal

Production by Examination of Degenerating Germ Cells. Russell and Clermont (1975) used electron microscopy to examine degenerating germ cells in the rat. One advantage of electron microscopy is that cells in the process of degeneration might be detected, whereas they would probably be included as normal cells if counts were obtained under light microscopy. Also, degenerating cells could provide an extremely sensitive evaluation procedure, because a very slight decrease in spermatogenesis might be accompanied by a several-fold increase in the number of degenerating cells (L. Russell, per- sonal communication). Obviously, the technique can become exceedingly time consuming, particularly if representative tissue sampling is to be insured. Huckins (1972) reported that degeneration of germ cells in the rat appeared to affect all cells joined by intercellular bridges. Thus, the importance of evaluating a testicular sample of adequate size is evident. This technique would appear to merit further evaluation for use in specialized studies, but is probably too difficult, time consuming, and expensive for use on a routine basis.

Accuracy of Procedures for Determining Absolute Rates of Spermatozoal Production. It is presently impossible to assess the true accuracy of procedures for estimating daily spermatozoal production. Gebauer et al. (1974a) reported that the daily spermatozoal output of stallions represented 81 to 98% of daily spermatozoal production estimated from quantitative testicular histology. Corresponding values based upon histological or homogenization techniques ranged from 63 (Kennelly and Foote, 1964) to 83 to 88% (Swierstra, 1968b) for the boar, 46 (Orgebin-Crist, 1968) to 82% (Holtz and Foote, 1972) for the rabbit, and 25 (Swierstra, 1966), 42 (Amann and Almquist, 1962) and 92% (Amann et al., 1974) for the bull. It is obvious that either spermatozoal losses through resorption in the excurrent ducts, micturition, or masturbation are exceedingly large, or spermatozoal production rates have been grossly overestimated in some studies.

Investigation of Precise Events in Spermatogenesis

Often it is desirable to determine if a reduction in spermatozoal production resulted from fzilure of specific cells to divide, or if greater cellular degeneration occurred, and, if so, which



830 BERNDTSON

cells u n d e r w e n t degenerat ion. The procedure by which this may be accompl ished is vir tual ly identical to tha t for s tudying relative changes in spermatozoa l p roduc t ion rates p roposed by C le rmon t and Morgenta ler (1955) in which one determines the mean number of germ cells per tubular cross sect ion. However , specific events are investigated by de termining the n u m b e r o f germ cells of each type in stages immedia te ly preceding and fo l lowing the event of interest , f rom which one can de termine the percentage of the expec ted cellular yield actually obtained. This procedure also has been ut i l ized to s tudy the kinetics of spermatogenesis and s tem cell renewal. While such in fo rmat ion is ex- t remely useful, greater insight into the s tem cell renewal process has been achieved with ad- dit ional techniques . Procedures for s tudying s tem cell renewal are beyond the scope of this review, and in teres ted persons are referred to the excel len t review by C le rmon t (1972) as well as references dealing with the examina t ion of spermatogonia l renewal in whole moun t s of seminiferous tubules (Cle rmont and Bustos- Obregon, 1968; Huckins and Kopriwa, 1969; Huckins, 1971a, b, c) for fur ther discussion of this subject.

CONCLUSIONS

The techniques described in this review evolved because of a need to object ively eval- uate the quant i ta t ive aspects of spermatogenesis. These procedures are imperfect . None the - less, when used judiciously, they are powerfu l tools for investigating the gametogenic act ivi ty of the testes, and should be ut i l ized whenever a critical assessment of the quant i ta t ive aspects of spermatogenesis is required.

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