The original purposes of this volume were to provide instructions for research-related duties and to standardize normative data collection procedures at the University of Washington Infant Primate Research Laboratory (IPRL). As such, the research manual and the technician's guide have been used to train students, new staff, and research investigators and their personnel since 1976. The material has been updated since that time as new standardized research protocols and husbandry techniques have been instituted. The documents are also used as appendix materials in support of applications for research grants and contracts.

As in the past, we are making the manual available to the scientific community at a charge that supports the cost of production and mailing. Our purpose is to provide researchers and colony managers with a model of a research-oriented nursery for nonhuman primates. The model illustrates the scope of the husbandry activities and types of normative developmental data collection pursued in the IPRL. These methods and data support the following major IPRL functions:

• Rearing of healthy neonates and infants in support of biobehavioral research projects.
• Preservation of at-risk newborns to serve as models of natural disease processes and developmental risk factors. These infants include low-birth-weight, premature, abused, neglected, and clinically ill neonates, as well as healthy neonates whose mothers cannot care for them due to illness or death.
• Generation of normative developmental data for husbandry purposes, for defining developmentally normal versus abnormal monkeys to determine their appropriateness as subjects in research projects, and for use as control data in experimental studies. The latter use has the goal of minimizing the numbers of nonhuman primate subjects required in many types of invasive research projects.

We believe the IPRL is unique in several ways. The Infant-Save program is universally applied in our Primate Center, and has preserved over 700 neonates and infants of three species that would otherwise have died or been euthanized in the interests of time and economy. Instead, the animals have been used as research subjects or have become members of the breeding colony. The collection of normative data is a formal core function, which supports technical personnel trained in the study of growth and behavioral development. These data are available without cost to researchers using the IPRL. Without such core support, most of this invaluable information would not be available, as few individual projects could support the cost and time required to generate these large data bases. A final aspect of uniqueness concerns the funding sources for the IPRL, which come from two separate centers supported by different institutes of the National Institutes of Health.

We would like to acknowledge the foresight of two individuals in making the IPRL and its functions possible—Orville Smith, former director of the Regional Primate Research Center (RPRC), and Irvin Emanuel, former director of the Child Development and Mental Retardation Center (CDMRC). As center directors in 1972, these individuals were responsible for proposing the establishment of the IPRL and funding it as a joint core facility of the two centers. The funding has come from NIH grant RR00166 from the Animal Resources Branch to the RPRC and grant HD02274 from the Mental Retardation branch of NICHD to the CDMRC. This support has continued over the last 20 years. During this time, the IPRL handled over 1200 nursery-reared infants and provided services, data, and husbandry and research training to over 200 university faculty researchers, postdoctoral researchers, and graduate students. Research training was also provided to 40-50 undergraduate students each year.

The materials in these documents, as well as many of the husbandry and research procedures, were generated by a number of IPRL technical staff members and graduate students over a 20-year period. We are indebted to them for the smooth functioning of the facility and the contents of this document. The major contributors include, chronologically, Jonathan Lewis, Ph.D.; Richard Holm, Ph.D.; Carol Fahrenbruch, MPH; Thomas Burbacher, Ph.D.; Sherry Savage, R.N.; Beth Goodlin-Jones, Ph.D.; Lauren Wasser, Ph.D.; and Coleen Walker, B.S.

We express our special thanks to Kate Elias, Primate Center editor, for her major contributions in designing, organizing, and editing this manual. Marjorie Domenowske, Primate Center illustrator, also merits appreciation for designing and drawing many of the illustrations.

Gerald C. Ruppenthal and Gene P. Sackett
Seattle, Washington
July 1992

[Table of Contents]

Blood Draws and Assays
Maternal Blood
Maternal blood draws consist of 5 ml taken at days 10, 13, 16, and 19 after timed mating for progesterone confirmation of pregnancy. Plasma samples must be frozen until assayed.

Blood is obtained on gestational days 30, 50, 70, 90, 100, 110, 120, 130, 150 and 165 and one week postpartum. Draws are done at the ultrasound examination with the animal under ketamine-rompum anesthesia, and just prior to a Cesarean section.

Draw 12 ml of whole blood by syringe: 7 ml on heparin for plasma assays (green top tube) and 5 ml for serum assays (red top tube). An addition 3 ml of blood for serum assays (red top tube) is drawn one week postpartum.

Do not centrifuge the blood. Place it upright on a test tube rack immediately upon drawing, keep it at room temperature, and take it to the laboratory within an hour of collection. All blood and amniotic fluid must be properly labeled (see example). Use adhesive labels and mark them with Sharpie marking pens. Do not write directly on the container.

Hematocrit and hemoglobin are determined from whole blood. Dams with hematocrits below 30 are given iron supplements. CBCs are done if dams display signs of illness. Other assays are shown in Table 1.

Plasma volume estimates are provided via Evans-blue dye technique on dams assigned to the 165-day C-section group. These females have an additional 12 ml of blood drawn for the Evans-blue procedure on days 40, 80, and 140 from conception, when they are anesthetized for ultrasound examination. Other blood draws are not scheduled for these animals. If the hematocrit of an otherwise eligible dam falls below 30, the Evans-blue procedure is skipped and iron supplementation is begun.

Fetal Blood
Fetal blood (> 2 ml) is collected on preservative-free heparin from all fetuses sacrificed at C-section. The blood is assayed as shown in Table 2. Surviving infants from the 165-day C-section group have only 2 ml of cord blood sampled.

Amniotic Fluid
Amniotic fluid is obtained from all dams at the 90-day ultrasound exam, and again at Cesarean section. The amniotic fluid (3-5 ml at 90 days and as much as possible at C-section) is put in plastic tubes, labeled as shown for blood samples, and assayed as shown in Table 3. The cells collected at 90 days are screened for Barr bodies by the histology lab for prompt sex determination. Spleen and skin tissue are used for karyotyping.

[Table of Contents]

Subjects
Pregnant female monkeys.

Equipment
22-gauge needles and butterflies; 1-cc, 3-cc, 5-cc and 10-cc syringes; sodium heparin; atropine; gauze pads; alcohol wipes; Vetalar; 5-cc heparinized vacutainers; 1-cc EDTA tubes; stopwatch; Doppler stethoscope; prepartum data sheets (Appendix, form 1); rectal probe and temperature box.

Time of Exam
Between 8 a.m. and 4 p.m.

Schedule
All exams are conducted on the same day unless an animal exhibits signs of fetal or maternal distress (see Guidelines for Preterm Termination of Pregnancy).

Procedure
All blood draws and procedures are conducted after the animal has been anesthetized with vetalar and atropine.

1. Fill a 3-cc syringe with 0.2 mg atropine (0.5 cc) and, at most, 10 mg/kg of Vetalar. Squeeze the animal up to the cage front and use a 22-gauge needle to inject the syringe contents into a large muscle site. The most common injection site is the large thigh muscle. Record time of injection on prepartum sheet.
2. Remove the animal from her cage and take her to the area where the examination table and supplies are located.
3. Weigh the animal and record her weight on the prepartum sheet.
4. Insert the rectal temperature probe and while waiting for it to stabilize, check for lactation. Record rectal temperature, time since vetalar injection, and yes/no for lactation.
5. With the animal positioned on her side, check fetal orientation. Roll her to a supine position to confirm orientation. Verify orientation by bimanual external palpation. Record the results on prepartum sheet (e.g., head down).
6. With the animal lying supine, prepare the femoral area for blood draw with an alcohol wipe. Draw 4 cc of blood with a 5-cc syringe and 22-gauge needle or with a vacutainer set-up. Occlude the femoral area with external pressure for several minutes after completing the blood draw to prevent any internal bleeding.
7. Divide the 4 cc of blood by unscrewing the needle, and with little pressure, put 3 cc into heparinized (green top) vacutainer and 1 cc into EDTA (purple top) vacutainer. Write the animal's number on both vacutainers for easy identification.
8. Roll the animal onto her side for fetal and placental heart rate measurements. Place lubricating jelly on her abdomen and use the Doppler stethoscope for counting fetal heart rate. Count for 15 sec and multiply by 4 to calculate beats per minute. Record fetal heart rate and placental rate on prepartum sheet. Ideally, two persons should count the beats simultaneously so that you have two counts to record.
9. Immediately after counting the fetal heart rate, count the maternal heart rate with the Doppler. Be suspicious of identical rates. You may have counted the maternal arterial rate rather than the fetal heart rate. Recount fetal heart rate if needed.
10. Maintain the animal on her side for the cervical palpation. With a clean glove and ample lubricating jelly on the fingers, slowly insert one finger into the vaginal canal. Place the other hand on the female's abdomen and apply slight pressure on the fetus so that the presenting part of the fetus moves toward the cervix. Slowly move the finger inside the vagina until the cervix is palpated. Determine the degree of effacement and dilation of the cervical ora, both the internal os and the external os. Record the state of the cervical ora on the prepartum sheet. Also, record whether you could palpate the fetus or not.
11. Check the animal's femoral area for any signs of internal bleeding. Clean off the lubricating jelly. Return her to her cage, laying her on her side to prevent saliva from collecting in her throat.
12. Clean the examination table and replace any supplies needed.
13. Use the large centrifuge to spin the blood samples. Spin the blood for a total of 10 min (two full cycles). Draw off the plasma with a glass pipette and rubber bulb and put it into a pink storage tube with a red snap-on top Use a ballpoint pen and label with a paper label sticker for the pink tube with the following:
    • animal number
    • date (mo/dy/yr)
    • study
    • prepartum
    • any other relevant information
    Place the completely labeled tube in a rack in a freezer for temporary storage.
14. Use a hematocrit centrifuge for hematocrit tubes. Put on gloves and then fill two hematocrit tubes with blood from the EDTA vacutainer. The hematocrit tubes should have some suction effect so filling the tubes should be easy. Close off one end of each hematocrit tube with the white caulk found with/near the hematocrit tubes. Place the hematocrit tubes into the hematocrit centrifuge for the maximum amount of time (+5 min). Once the centrifuge has stopped, read the hematocrit tubes on the reading calibration machine, or use the manual reading card if necessary. Discard the tubes and the remaining sample in the EDTA tube once you have results that are identical or within 2 points of each other. If the two samples differ by more than 2 points, redo the entire process with another sample of the blood from the EDTA tube. Record the hematocrit (or PCV) values on the prepartum sheet.
15. Make any needed comments about any female that is close to delivering, noting each female's cervical status, etc. Place comments where staff members can see them.
16. Place a note stating that an animal has been anesthetized with a note on the time of anesthesia, so that status checkers know not to code her behavior.

[Table of Contents]

The Purpose of Status Data

• The primary reason for collecting data on the status of pregnant females is to study changes in their activity patterns as delivery approaches. Of interest is the way in which individual patterns of activity may be related to any concurrent or previous experimental condition (i.e., reproductive history, exposure to toxins, etc.), pregnancy outcome, and infant characteristics such as sex, survivorship, and birth-weight centile.
• Another reason for collecting status data is to detect signs of labor. Some indications of labor are frequent postural changes, stretching, and straining against the side of the cage. Any female that is awake and active after 7 p.m. should be watched closely. If you decide that a female is in labor, start the videotape to record her behavior and contact the appropriate delivery team personnel.
• A third important reason for taking status checks is to monitor the physical condition of each animal. Animals may experience illness or infection during their stay. Signs of potentially dangerous health problems include vomiting, lethargy, and weak or wobbly movements. If you notice any unusual behaviors that you think could indicate a female is in trouble, do not hesitate to call it to the attention of the veterinary staff. It is better to have a false alarm than a delay in treatment.

Procedure
The status of each pregnant female is recorded on data sheets (Appendix, form 2) every 30 min. Animals are monitored by video via closed-circuit TV. If you are unable to get a satisfactory picture on the television monitor, try adjusting its brightness and contrast controls.

Maternal Status Codes

1—ASLEEP: May be sitting or reclining, eyes closed. If her back is to the camera, watch her arms for movement.

2—PASSIVE: Awake but not doing anything. This code includes dozing and non-attentional looking (i.e., eyes open but not focused on anything).

3—QUIET ALERT: Awake and doing something (i.e., grooming, cage licking, attentional looking). No locomotion.

4—STANDING OR WALKING: Either standing still or slow walking.

5—VERY ACTIVE: Very active locomotor behaviors (i.e., pacing, backflips, cage shaking, locomotor stereotypies).

6—EATING: Self-explanatory.

7—DRINKING: Self-explanatory (the drinking bottle or "Lixit" is located on the side of the cage).

8—VAGINAL EXPLORE: Fingering the vaginal area. (Grooming the ventrum is not a vaginal explore!)

9—BIRTH: Although birth is rarely seen during status checks, the event is still coded in the half-hour time line closest to it.

On Becoming a Status Observer for Females and Infants

1. Talk and walk through the procedure with the supervisor. Review definitions, location of data sheets, and other instructions.
2. Practice coding adult females on the training tape.
3. Practice coding nursery infants with trained staff.
4. Take reliability test on videotape. Must reach 90% agreement (18 out of 20).
5. Repeat steps 1-4 if needed.

[Table of Contents]

One of the most important responsibilities of the nursery staff is to observe pregnant females for signs of labor. A tremendous amount of research effort and money goes into each pregnancy, and by the time a female arrives at the lab, a significant amount of data has already been collected. The birth data are an integral part of the research design. They serve as an endpoint for fetal information and are a basis for later testing and observations.

Observers
Designated technician or animal care personnel, especially night workers, as most deliveries occur between 8 p.m. and 1 a.m.

Procedures
Monitor each female as often as you can, but at least once every 30 minutes. Spend enough time at each status check to see if she is showing signs of labor. When in the nursery, keep an eye on the monitor in there. Signs to watch for are:

• digital exploration of the perineal area, followed by sniffing and licking the fingers
• restlessness, indicated by shifting postures to get comfortable
• pacing
• cyclic activity such as drinking, walking around, lying down, and repeating the pattern
• hanging on the cage and pushing down
• postures unique to labor

Sometimes you can see contractions. Start the videotape at the first sign of labor. You can always record over a false alarm if the dam settles down and goes back to sleep. There are several sample tapes of labor and deliveries. To see one, make an appointment with the research technologist in charge.

Reference
Goodlin, B.L. and Sackett, G.P. Parturition in Macaca nemestrina. Amer. J. Primatol. 4:283-307, 1983.

Tester
On-Call Delivery Teams: check telephone listing for individual on duty call

Equipment
Sterile birth pack, 1-cc and 5-cc syringes, Vetalar, 5-ml green-top tubes, 1-ml purple-top tubes, small-animal respirator, diapers, birth record forms (Appendix, forms 3 and 4), microhematocrit tubes, microhematocrit centrifuge, centrifuge, distilled water, blood draw clipboard, stopwatch, incubator, microhematocrit wax, rectal thermometer, local or other disinfectant, plasma storage tubes, plasma storage tube labels, pipette, bulb for pipette, heat seal, heat seal bags, sterile petri dishes.

Time of Test
At birth

Pre-Delivery
Nursery Staff
You are responsible for recognizing labor behavior. Familiarize yourself with the females that are expected to deliver when you begin your shift. Watch the composite labor tape to learn what labor behavior looks like. Be observant while collecting the status checks on the females. If a female is awake but not active and is near her term-date, tape record her behavior. Continue to move the camera away from her to collect status checks on other females but always reposition the camera on her when you are through. Check her behavior every 10 minutes to see if she has begun to labor.

Delivery
Nursery Staff
Whenever you think a female is in labor, start the videotape recorder and fill out a videotape form (Appendix, form 3). Then call the appropriate delivery team person, who may be using a bellboy number. If you are unable to reach the person, call the next person listed on the on-call schedule. Keep calling until you reach someone who will come in to the lab. Until the delivery person arrives, continue the videotaping; start new tapes if necessary.

As soon as the delivery person is in the lab, the videotaping and all of the delivery procedures are his or her responsibility. When the delivery person arrives, tell him/her about the videotaping, which mother is in labor, and any other information.

Procedures
Delivery Staff
Upon arrival in the lab, find out which animal is in labor, when labor began, and whether videotaping is in progress. You are responsible for maintaining the videotaping until delivery. Get the data papers needed to record information (forms 3 and 4 in Appendix). Fill in any information that you can while waiting for the delivery.

Suggested supplies to maintain are:

• Incubator and distilled water and cloth cover
• Placenta exam equipment
• Delivery pack (clamp, scissors, bulb syringe)
• Rectal temperature probe and reading box
• Blood sample supplies
• Vetalar, atropine, heparin
• Stethoscope
• Stopwatch
• Syringe with black-top needle, with correct amount of vetalar and atropine for the dam

Post-Delivery

1. Note the exact time of delivery. Write it down on the data sheet and start the stopwatch.
2. Administer vetalar/atropine to female. Note time of injection on data sheet.
3. Put blue pads on examination table; fill a 5-cc black-top syringe with heparin.
4. Check incubator temperature. Turn up to 93° if needed. Put distilled water in incubator basin.
5. When the dam is sedated, roll her on her back, place the infant on her chest, and carry both to the examination table.
6. Insert temperature probe into female's rectum.
7. Check the infant's breathing. Use bulb syringe to clear mucous from nostrils and mouth. Count infant's respiration and heart rate. Record dam's temperature, wipe probe with alcohol wipe, and insert into infant's rectum. Record infant's first Apgar score—respiration, heart rate, and temperature—on birth record (form 4 in Appendix) and on infant evaluation sheet (form 5). Note time since delivery.
8. Open delivery pack and clamp the umbilical cord about two inches away from the infant's abdomen. Get 4 hematocrit tubes within reach. Cut the umbilical cord and hold the hematocrit tubes up to it. Allow the umbilical blood to fill the tubes and then close off the tubes at one end by pressing an end into the white caulk strip.
9. Place infant in incubator and rub it with diapers.
10. Take a blood sample from the female's femoral vein. Occlude the vein for at least 60 sec.
11. Take the second set of Apgar measurements on the infant. Take respiration first, then heart rate, then temperature. Note time since delivery.
12. Check the female for lactation. Record on labor and delivery data sheet (form 3 in Appendix).
13. If the placenta hasn't been delivered yet, massage the female's abdomen and apply light pressure to the area at the top of her uterus. This should be enough manipulation to expel the placenta. If not, try again. If necessary, keep waiting for delivery of the placenta until the female begins to move. If she continues to retain the placenta, note it on the yellow placenta sheet and leave a note for the nursery staff. Ask them to watch for its delivery and to note the time.
14. Weigh the female. Record on data sheet and placenta sheet. Put her back into her cage, positioned on her side so that saliva will not collect in her throat.
15. If the infant is scheduled for a perinatal blood draw, do it now.
16. Take infant to the nursery and put it in an incubator. Cover the incubator with the cloth cover and plug it in. Add more distilled water if necessary. If the infant has labored or difficult respiration, place it on oxygen.
17. Conduct the placental exam as detailed in the following section. Wear a surgical mask while conducting the exam. After completing the exam, cut two cubes out of the center of each lobe and place each individually into a small vial. Completely label each vial with dam's number, date, and from which lobe the sample is taken. Place the remaining placental tissue in a large jar containing formalin. Label the jar with dam's number, date, and contents. Leave jar and vials in exam cabinet (see placenta exam instructions).
18. Conduct the remaining parts of the infant exam. Check infant's reflexes and record on the infant evaluation sheet (Appendix, form 5). Check all external features by following the list on the sheet. Do not do the anthropometrics. Head molding refers to the shape. Note any flattened or pointed areas. A fontanel may occur anywhere on the skull. It is a soft spot from incomplete ossification. Overriding sutures can be found along the suture lines when one of the skull plates protrudes above the other plate. Sternal retraction occurs when the sternum appears to fold in and out due to difficult breathing. When completed, leave the assessment sheet under the infant's green diurnal sheet on its clipboard.
19. Weigh the infant on the nursery scale and record the birth weight on the birth record form.
20. Centrifuge any hematocrit tubes in the small hematocrit centrifuge. Centrifuge any blood samples in the large centrifuge. Read the hematocrit tubes on the hematocrit reader. Discard them and record the values. Draw off the clear serum from centrifuged samples and put it in a storage tube. Label the sample completely. Put it in a tray in the freezer.
21. Fill out the rest of the birth record form.
22. Ask the nursery staff to measure the infant's temperature once an hour until it reaches 98.5°F.

If there are any questions or problems with the videotaping, measuring, etc., contact the supervisor.

It is best to do this exam close to the time of delivery, so do as much as you can at that time. If you cannot examine the placenta or are uncertain of something, place it in a plastic bag, refrigerate it (do not freeze it!), and leave a note so that someone can look at it the next day.

Supplies: scale, jars of formalin, plastic bags, heat sealer, calipers, scissors from the delivery pack, and placental examination form (Appendix, form 6).

Remember to wear a mask while doing the exam to avoid inhaling flakes of dried blood that may be floating in the air.

Placental Characteristics

Insertion of umbilical vessels
The umbilical cord is usually in the central area of the placenta. However, it may insert eccentrically (off center), or marginally (toward the edge).

Number of umbilical vessels
There are normally three umbilical vessels: two arteries and one vein. After checking the fetal end (the already cut end), cut the cord off about an inch above the placenta to check the placental or maternal end. If the three vessels are not apparent, it may help to squeeze the cord a bit so that a small amount of blood appears at the end of each vessel.

Number of lobes
The typical macaque placenta consists of two lobes. The umbilical cord inserts into the primary or "A" lobe and then gives off a number of vessels which run within the fetal membranes to supply the secondary or "B" lobe. Lobe A is usually larger than lobe B. If there appears to be only one lobe, check to be sure that there are not two lobes that have fused together.

Placental Abnormalities

Placental lobes fused
In some cases the two placental lobes may be joined at their margins. If you find a single-lobed placenta, always check to be sure it is not actually a case of lobe fusion. A slight line, depression, or crease of fetal membrane occurs between the lobes at the point of fusion. Fusion may be partial or complete. Complete fusion means that the lobes are joined for the entire length of one side. If the length of attachment is less than that, the fusion is partial.

Membranes meconium stained
Meconium is a brown-green-colored substance that is passed into the amniotic fluid by a distressed fetus. It may wash away when the fetal sac breaks during delivery and may not be detectable.

Placenta praevia
Implantation and development of the placenta in the lower uterine segment that is either covering or adjoining the internal cervical os (opening) is called placenta praevia. It may be complete, partial, or marginal. Because the placenta blocks the cervical opening, placenta praevia usually causes severe hemorrhage and possibly the death of the mother and/or fetus during parturition. C-section is usually required for a successful delivery. Placenta praevia may give the placenta a somewhat cone-shaped appearance because of its unusual uterine attachment, but is otherwise undetectable after delivery and probably hasn't occurred if the delivery was normal.

Insertion of membranes
The fetal side of the placenta is covered by the fetal membranes. This covering is called the chorionic plate. As the membranes leave the placental surface, they extend around the fetus, forming the fetal or amniotic sac. In some cases, the membranes become folded back upon themselves. Degenerated decidua and fibrin accumulate in the fold, creating a whitish ring of infarct on the fetal side of the placenta. This abnormality, called extrachorialis, may take either or both of two forms: circumvallate and circummarginate. In a circumvallate placenta the ring is composed of a double fold of amnion and chorion (membranes) with degenerated decidua and fibrin between them. In a marginate placenta, the ring is coincident with the placental margin, where the chorion and amnion are raised by interposed decidua and fibrin without much folding of the membranes, i.e., a margin of elevated infarcted tissue. Extrachorialis may be complete circumvallate or marginate, or any combination of circumvallate, marginate and normal.

Retroplacental hemorrhage
Retroplacental hemorrhage occurs when the placenta separates prematurely from its uterine attachment. The hemorrhage consists of clotted blood on the maternal (non-membrane-covered) side of the placenta. Both recent and old clots may be present. The older clots will be darker in color—dark to blackish red. Recent clots are light red (more orange) and may be associated with delivery. Note the age, size and extent of the clotting.

Retromembranous hemorrhage
Retromembranous hemorrhage is bleeding beneath the fetal membranes; it usually indicates marginal sinus rupture. Look for it on the fetal (membrane-covered) side of the placenta. Again, note the age, size and extent.

Infarcts
Infarcts are areas of tissue necrosis usually caused by loss or insufficiency of circulatory blood supply. They appear as areas of whitish, yellow or pink hardened tissue. Infarcts can be superficial or can extend through the entire placental thickness. Cut into the placenta at the infarct to see if it is deep or superficial. Often you will see extensive but superficial infarction covering the maternal surface. Deeper, globular areas of infarct may also occur, especially on the placental margin. You may want to wash off the placenta before looking for infarcts.

Placental Dimensions
Since placental dimensions are taken with the membranes and cord removed, these measurements should be taken last. Remove the cord (if you haven't already), and trim away the membranes. Lay the placenta flat on the table and move the calipers around the margin, noting the largest and smallest diameters.

Reference
Fahrenbruch, C.E., Burbacher, T.M., and Sackett, G.P. Gross placental morphology and pregnancy outcome in Macaca nemestrina . In G.C. Ruppenthal and D.J. Reese (eds.), Nursery Care of Nonhuman Primates. New York: Plenum, 1979, pp. 27-34.

All pregnant females have weekly physical examinations during the third trimester of pregnancy (+130 days). The examinations are conducted by a veterinary technician and/or research technologist at the beginning of each week until delivery occurs.

Indications that a pregnancy should be terminated before term include the following:

Prolonged, unproductive labor
Active labor, with physical straining and an intercontraction interval of less than 10 minutes, which continues for at least 30 minutes and then ceases prior to delivery.
Action taken: If the female's behavior is unremarkable and she does not appear to be in pain and is not bleeding from the perineal area, notify the research technologist or the veterinary technician to examine the female the next morning. The female will be allowed two nights of unproductive labor before a Cesarean delivery is performed. If at any time, the female appears to be in shock, is bleeding, or is behaving abnormally, call the veterinary technician or the research technologist for an immediate examination. They will arrange for an immediate Cesarean delivery if either the fetus or the female is in a compromised condition.

Prepartum bleeding or hemorrhage
Bleeding from the perineal area.
Action taken: Notify the research technologist or the veterinary technician to examine the female as soon as possible. A Cesarean delivery will be arranged if the female or fetus is in a compromised condition. If not, the situation will be monitored for 24 hours and, if the bleeding persists, a Cesarean delivery will be performed.

Placenta praevia
The placenta is attached to the uterine wall over the cervical os, as determined by manual palpation.

Action taken: Cesarean delivery prior to term.

Fetal death
No indication of body movements and absence of a fetal heart rate or placental rate.
Action taken: Immediate Cesarean delivery.

Maternal disease
Any acute or chronic disease of the female.
Action taken: Close monitoring of the female and fetus with frequent physical examinations to assess condition. If recommended by the veterinary staff, Cesarean delivery prior to term.

Contact the research technologist or veterinarian when considering a female's condition and these guidelines.

Reference
Mahoney, C.J. Clinical indications for cesarean section in the rhesus monkey. In G.C. Ruppenthal and D.J. Reese (eds.), Nursery Care of Nonhuman Primates, 1979, pp. 3-20.

[Table of Contents]

Subjects
Nonhuman primate infants

Testers
Research Technologists

Equipment
3-cc syringes, sodium heparin, 2-ml conical tubes, 21- or 23-gauge needles, Alcowipes, 4 x 4 gauze sponges

Time of Test
Between 4 and 6 p.m.

Schedule
At birth and at 2, 4, and 8 months

Procedure
At Birth
Draw 1 cc of blood from the femoral- vein into heparinized syringe. Transfer to 2-cc conical tube and label:

• "Infant of," dam's number
• Birth draw
• Date

At 2, 4 and 8 Months
Draw 2 cc of blood from the femoral vein into heparinized syringe. Transfer to 2-cc conical tube and label:

• Subject number
• Date
• Age (months)

Spin blood for 20 min at high speed, pipette off plasma, label, and freeze.

[Table of Contents]

Tooth Eruption

Subjects
Nonhuman primate infants.

Time of Test
Whenever convenient (good to do it while another task is being done on the animal).

Schedule
Daily from birth until eruption of all five baby teeth on both sides in upper and lower jaw.

Procedure
Examine each animal for the eruption of teeth through the gum. When the tooth has cut through the gum, record the date on the daily tooth check sheet (form 7 in Appendix).

Definitions of teeth are as follows: A—central incisors B—lateral incisors C—canines D—first molar E—second molar   The teeth usually erupt in that order. However, in some individuals, especially in long-tailed macaques, the first molars erupt before the canines.

Additional Protocol to Determine Degree of Eruption

1. Check every day for new monkeys in the nursery. If a new infant has arrived, start a new data sheet for it (form 8 in Appendix). Fill in the information at the top right-hand corner and write the day's date in the space on the left, with "f.c." (first check). If the incisors have already erupted, follow instruction #3 below. Follow the date with a "-?" because you don't know how many days ago the tooth erupted.
2. When a tooth is first observed to reach a certain stage, place the day's date for that tooth on the data sheet.
3. If a tooth has passed a stage by the time you see it, write in today's date for the highest stage it has reached and add a note of the number of days missed since the last time this tooth was observed. For example: A canine was at Stage 1 the last time you saw it, but you missed a week and the next time you saw it, it was already at Stage 3. Put a dash for the date of the stage you missed (Stage 2). For Stage 3, put the date of actual observation and write next to it "-7" for the missing week.

    

Use the following descriptions to determine the three stages of eruption in infant macaques.

Stage 1

Incisors, canines, and molars (Fig. 1):

If you observe any of the following in the location where the tooth you are observing is due to erupt, the tooth has reached Stage 1:

Elevated ridge or swelling Redness, and sometimes bleeding if rubbed
Fig. 1—Stage 1

Stage 2

If you observe any of the following in the location where the tooth you are observing has erupted, the tooth has reached Stage 2:

Incisors (Fig. 2): The tooth is visible—i.e., it has broken through the gum but the entire top surface of the crown is not yet through.	Fig. 2—Stage 2 incisors
Canines (Fig. 3): The tooth is visible—i.e., has broken through the gum, but has not yet reached Stage 3. There is a little white dot in the location where the cusp is about to break through. The tooth is not yet visible, but you can feel a sharp edge in the location where you expect the tooth to erupt.	Fig. 3—Stage 2 canines
Molars (Fig. 4): The tooth is visible, but only one or two cusps are showing. A little white dot is visible in the location where the first cusp is about to break through. The tooth is not visible, but you can feel a sharp edge in the location where the tooth is expected to erupt.	Fig. 4—Stage 2 molars

Stage 3

If you observe the following, the tooth has reached Stage 3:

Incisors (Fig. 5): The entire top surface of the crown has just emerged. (If more than just the top of the crown has emerged, it is beyond Stage 3.)	Fig. 5--Stage 3 incisors
Canines (Fig. 6): The cusp of the canine is level with the adjacent lateral incisor.	Fig. 6--Stage 3 canines
Molars (Fig. 7): All four cusps have erupted and the entire top surface of the crown has just emerged. (If more than just the top surface of the crown has emerged, it is beyond Stage 3.)	Fig. 7--Stage 3 molars

Anthropometrics and Radiographs

Objective
Anthropometric measurements of Macaca nemestrina infants are made at regular intervals up to a postconception age of 355 days. X-rays are taken to determine the degree of ossification of the epiphyses and round bones of the left hand and foot as a measurement of development.

Schedule
Animals are assessed at birth and at 173, 187, 215, 245, 271, 299, and 355 days postconception; a ±2-day margin is acceptable. Consult the calendar in the "Anthropometrics" folder to see which animals are schedule to be measured and radiographed. On the anthropometric and ossification data sheets (Appendix, forms 9-11), fill in the date and the animal's age. If assessment is being done early or late, note the scheduled date and the animal's age.

Equipment
Calipers, sliding scale (Fig. 1), scissors, tailor's tape (1/2 inch width), head circumference measuring tape (Fig. 2), x-ray film, lead gloves, x-ray viewing machine, magnifying glass, and anthropometric and ossification data sheets.

Fig. 1—Sliding scale	Fig. 2—Head circumference measuring tape

Procedures
Anthropometrics
Each measurement is made by two persons. Do not disclose individual measurements until both have been taken. Interobserver measurements that vary by more than 3% must be repeated. Measure head circumference, length, and width to the nearest millimeter; measure foot length and crown-rump length to the nearest tenth of a millimeter. Record the two scores and their average on the data sheet. If the average is not an integral number, round up at > 0.5 (e.g., 233.5 becomes 234).

Head circumference: Measuring tape should encircle the head below the brow ridge (covering the animal's eyes), along the nuchal crest above the ears, and should cover the occiput. Pull the tape tightly through the clamp and then fasten clamp at the side of the head. Read the measurement while the tape is on the animal's head.

Head length (Fig. 3). With the spreading calipers, measure the length from the area between the eyes to the occiput. If the animal is disturbed by the procedure, blindfold it with tailor's tape. Read the measurement while the calipers are in place.

Fig. 3—Head length

Head width (Fig. 4). With the spreading calipers, measure between the two widest points of the skull; the landmarks are just above and in front of the ears. Feel for these points first and then replace your fingers with the calipers.

Fig. 4—Head width (Frankfurt plane)

Left foot length (Fig. 5). Place the monkey's heel flush with the base of the sliding scale. Center the foot directly over the scale, with toes aligned and completely extended. Read the length from the end of the longest toe, excluding the nail. For accuracy in reading, you must be positioned directly over the scale.

Fig. 5—Measuring foot length

Crown-rump length. If the animal is wrapped in a diaper, move the diaper away from the dorsal surface so that it does not come between the animal and the sliding scale. Allow the animal to clutch the diaper ventrally as this procedure is usually stressful. Place the animal on the scale in a supine position with its rump against the base of the scale and its tail to the side. Hold the animal's feet and lift its legs to facilitate positioning of the rump. Apply pressure to the ventral trunk so that the animal does not arch its spine. Hold the head steady in the Frankfurt plane (see Fig. 4). Bring the sliding mechanism down to the crown. Holding the sliding mechanism firmly in place, lift the animal away from the scale and read the value while positioned directly above the scale.

Weight. Put the infant in the weighing bowl on the scale. Allow it to clutch a diaper for security. To adjust for the weight of the diaper, place a separate diaper in the weighing bowl and push "Tare" button. Remove the diaper and place the animal with the security diaper in the bowl. As animals rarely remain stationary during this procedure, estimate the weight as that figure which appears most consistently on the digital display. Check this weight against that taken daily by the technicians in the Infant Lab.

Radiographs
Radiographs of the animal's left hand and arm and left foot and lower leg are to be performed at the time of each anthropometric assessment. Take the animal to the x-ray suite in a clean transport cooler. Be certain that the locking mechanism is working. Coolers without the "Live Animal" label are preferable because they attract less attention in the hallways. Notify lab office personnel which animals are being taken to x-ray. Bring the animal's anthropometric data sheets, extra diapers, several pairs of surgical gloves, and x-ray film. Use Kodak Diagnostic Film, X-OMAT AR, 50/35 x 43 cm (cat. #165 1678). One sheet of film is sufficient for two animals, but bring extra sheets in case x-rays need to be redone.

Loading film. In the darkroom, turn on the developer and close the lid. Lock the door, turn off the main light, and turn on the infra-red light. Remove film from the envelope, taking care to separate film from paper sheets. Do not discard envelope. Insert film in the cassette and place the cassette in the door marked "unexposed," which is connected to the x-ray room.

Taking x-rays. The x-ray machine should be set at 200 milliamperes, 1/30 seconds, 42 peak kilovolts. The light source should be placed one meter above the cassette; check this distance at each radiography session. Place the cassette on the x-ray table and cover half of the cassette with lead shields.

Place the animal on a folded diaper at the edge of the cassette and cover its body with lead gloves. Extend the animal's left leg out onto the cassette. Use gauze tape around the knee and elbow joints before taping the limbs to decrease discomfort to the animal. Tape the left leg down at the knee joint and the foot. Extend the left foot as far as possible and hold the hallux at a 90° angle away from the toes. Such placement decreases the amount of superpositioning of the tibia and fibula, thereby maximizing the clarity of the areas of interest. Tape the left arm down at the elbow joint. See that the fingers of the left hand are aligned, flattened, and taped with the pollex set off from the other digits. See Fig. 6 for proper placement.

Fig. 6—Placement for x-ray

When finished, replace the animal in the cooler and clean the cassette, table and lead gloves with disinfectant. Cover the exposed side with lead shields and use the unexposed side to x-ray the next animal. Place the exposed cassette in the door marked "exposed."

Developing film. With the door locked and infra-red light on, remove the film from the cassette and feed it into the x-ray machine. Wait for the second safety light to appear before turning on the overhead light. X-rays take approximately 3-4 minutes to develop. Check the developed x-rays for clarity and detail (i.e., lack of superpositioning, clarity of carpals and tarsals). Return the x-ray to the original envelope for transport. When you are finished, be certain that both the x-ray machine and developer are turned off.

Concluding steps. (1) Return the animals to their cages in the nursery or quad room. (2) Disinfect the coolers. (3) Return the data sheets to the folder and cross off the animal numbers on the calendar.

Reading radiographs. The radiographs reveal the state of ossification for the epiphyses of the metacarpals, metatarsals, and phalanges, the carpals and tarsals, and the distal epiphyses of the ulna, radius, fibula, and tibia. To read the radiographs, you will need a magnifying glass, an x-ray viewing machine, hand and foot ossification data sheets, and diagrams of the bones in the hand and foot (Fig.7).

There are three possible ossification scores for the epiphyses and round bones:

• Absent (score 0.0)—There is no visible bone present.
• Initial (score 1.0)—Beginning ossification of the bone is barely visible. This is the most difficult stage to identify because it appears as a smudge on the x-ray. It looks like a small, cloudy white spot with no definition around the edges.
• Present (score 2.0)—Ossification of the bone is clearly visible. The bone may be small, but it has a definite shape and sharper white appearance than the initial stage of ossification.

Two scores for each hand and foot are tallied and entered into the Bone Data File. The foot score (fs) and hand score (hs) are computed by adding the total score from the ossification data sheet. The foot number (fn) and hand number (hn) are the actual number of centers present (i.e., any center with the score 1 or 2).

References
Newell-Morris, L., Tarrant, L.H., Fahrenbruch, C., Burbacher, T.M., and Sackett, G.P. Ossification in the hand and foot of the pigtail macaque. II. Order of appearance of centers and variability in sequence. Am. J. Phys. Anthrop. 53:423-439, 1980.

Newell-Morris, L., Carrol, B., Covey, A., Medley, S., and Sackett, G.P. Postnatal growth and skeletal maturation of experimental preterm macaques (Macaca nemestrina). J. Med. Primatol. 20:17-22, 1991.