A study of some factors influencing fertility and sterility in the bull

by Herbert Lester Gilman

Technique

The material used in the work came from abattoir animals, bull calves and adult bulls raised in the department herd, and sires upon which clinical observations had been made by various veterinarians in the field. Semen samples, many of which were sent in, were collected as often as possible after the method described by Williams (16). The genital organs were removed with as little chance of contamination as possible, and taken or sent to the laboratory where the examinations were made soon after arrival.

All cultures were made by searing the surface carefully, tearing out a small portion of the tissue with sterile forceps, and placing it upon the media. In most cases, however, where fluids were present, tubes were inoculated with the material which had been drawn off with a sterile pipette. As stated by Carpenter (9), in his work on the female genital tract, the organisms usually live in the depths of the tissue. The media used principally were glucose glycerin agar (glucose 1 per cent, glycerin 3 per cent); plain agar, both with a pH value of 7.4, and Loeffler’s blood serum. Small amounts of sterile blood serum or defibrinated blood were added to most of the agar slants to insure better growths of streptococci when present. All tubes, to which the serum had been added, were incubated for forty-eight hours before inoculation to insure absolute sterility.

After inoculation, the agar tubes were sealed with sealing wax to give a partial oxygen tension which was quite necessary in isolating the streptococci. The growth of other organisms was by no means hindered by the procedure, for one tube from each organ was often left unsealed. Incubation was at 37° C, and the routine method of examining the tubes was identical with the method of Carpenter (9).

Whenever possible, a sample of blood was obtained from the animal either before, or at the time of slaughter, for agglutination with Bact. abortum antigen. Extracts from the seminal vesicles, testes, and epididymes were injected into the guinea pigs and examined at the end of four to six weeks for the presence of Bact. abortum.

Sections of all organs were fixed as soon as possible in either Zenker’s or Helly’s fluid. Hematoxylin and eosin were used as routine tissue stains. Eosin and methylene blue, and Mallory’s connective tissue stain were, however, frequently utilized for special staining reactions.

The motility of the spermatozoa is best observed about half an hour after ejaculation, when the thick tenacious clot has started to liquefy. A drop of the fluid is placed upon a warmed slide, preferably one with a slight depression in it, and observation made with high or low powered objectives. The semen may be examined whole, or diluted with physiological saline solution. In the latter case, the sperms have a greater opportunity for freedom of motion in the absence of the thick viscid coagulate. A small vial of saline solution may be carried in one’s pocket where it will be kept warm, and a drop of this placed upon the glass slide. If the clot of semen is merely touched to this drop on the slide, plenty of spermatozoa will be deposited for an examination. This method is very satisfactory for the observation of motility, but needless to say, the undiluted semen must be used for the determination of the number of sperms present. If necessary, the specimen may be covered with a cover glass and the oil immersion objective used. While a warmed slide is quite sufficient to enable one to detect the presence of motility, the field soon cools and the sperms gradually become less motile. If possible, it is best to use a small electrically heated stage warmer, which keeps the field at a constant body temperature, so that the duration of the motion may be observed for hours if warmed physiological saline solution is added as the fluid evaporates.

Stained preparations are best made with thin smears on the glass slides. This is conveniently done by placing a drop of the semen on a slide and smearing it over the surface with the edge of another slide. A fairly thin and even field is thus obtained. A still better method is to first dilute the semen with physiological saline solution, so as to obtain fewer sperms in the field. After drying the preparation in air, fixation may be produced by drawing the slide through a gas flame several times, by immersion in equal parts of alcohol and ether, or even by the use of tissue fixers such as Helly’s or Zenker’s fluids. For ordinary staining, heat fixation is the quickest, and at the same time is quite satisfactory. After the slide is cooled, or washed, to remove the fixing solutions, it should be placed for a few minutes in a freshly prepared solution (1 per cent) of chlorazene, as recommended by Williams, to remove the mucus and proteid material which otherwise blur the field. Other authors (38) have recommended diluting the semen with about twenty volumes of a 0.12 per cent sodium carbonate solution in 0.8 per cent sodium chloride. From this liquid the cells should be centrifuged for several minutes, removed with a pipette, and smeared on the slide. Following this, the slide should be thoroughly washed, preferably for ten minutes in running water, after which it is ready for staining. Numerous methods have been used for this procedure, but the sperms are more or less erratic in their reactions to the dyes, and one must be very careful to use the same method in all samples, in order to obtain uniform results. For quick staining, to bring out gross abnormalities of structure, and number of sperms present, one may use Gram’s stain, or a light stain with any of the aniline dyes, such as fuchsin. To bring out the finer structure, particularly of the head, more careful technic must be employed.

Carnett and others (38) recommend the following: “The method of staining by iron-hematoxylin, particularly when supplemented by a cytoplasm stain, has proved, on the whole, the most satisfactory, and possesses the additional advantage of being absolutely permanent, a quality that few anilines can boast. The method consisted of treating the fixed object—and here the fixing agent was heat—with a two per cent solution of iron-alum for from two to four hours. The excess of iron-alum was then completely removed by pure water, and the object treated with a solution of hematoxylin (one per cent aqueous) for twelve hours or longer. The cells by this time were perfectly black. However, a 1 per cent solution of iron-alum removed the stain from the cytoplasm, leaving the chromatin of the head, the centrosome, and the axial filament a brilliant blue-black. Care must be taken that the preparation is not over-decolorized. After decolorization a saturated aqueous solution of eosin was added for from one to three minutes. This stained the protoplasmic envelope pink, and, unless the envelope is overstained, the view of the inner structures is not impaired in the least.”

Williams (17) recommends using two staining solutions, one of alcoholic eosin and fuchsin, the other a diluted methylene blue. The results obtained are, however, more or less erratic, due to the unstable character of the former stain, and the ease with which one may over or under stain. Many beautiful specimens may, nevertheless, be obtained by this method. I have frequently used a fairly quick method, though one not satisfactory in all cases, which consists of staining for five or six minutes in a saturated aqueous solution of fuchsin, washing in water, and counterstaining for a few seconds in a strong solution of methylene blue. A quite satisfactory method is to stain from two to five minutes in a saturated aqueous solution of methyl green, with the application of gentle heat. The heat may be applied by warming the slide over a gas flame as it steams, or by placing the jar containing the stain in a hot water bath. The slide is then washed thoroughly and counterstained for five minutes in a strong aqueous solution of eosin. This is a fairly reliable method, and many excellent preparations may be obtained by its use. The nucleus is stained green, the anterior part of the head and all of the tail pink. So far, I have found this a very reliable stain for routine work.