Sites for venous blood sampling include the jugular vein (Fig. 7-1), cephalic vein, lateral thoracic vein, saphenous vein, and coccygeal vein. Any vein that can be identified, occluded, and accessed safely may be used. The technician should use an approach and preparation similar to that for an intravenous (IV) injection, described under Parental Injection Techniques.

FIGURE 7-1 A, Location of the jugular vein B, Use of a Vacutainer for blood collection from the jugular vein.

Sites for arterial blood sampling are more difficult to access, especially in unsedated patients. The transverse facial artery and the dorsal metatarsal artery are most commonly used, especially in anesthetized patients. Arteries are prone to development of hematomas after needle withdrawal, so firm pressure should be applied directly to the puncture site (minimum 1–2 minutes) to minimize this risk.

Blood can be collected by aspiration into a syringe or Vacutainer tube. If dealing with a foal or needle-shy adult, Vacutainer tubes can be challenging to use because of patient motion. When patients are needle-shy, it may be advisable to use a 20-gauge (ga) needle and Luer syringe to aspirate the blood, then transfer it to a vacuum tube. For direct blood draws into Vacutainer tubes, -inch-long Vacutainer needles are available for large animal use.

Processing and analysis of blood samples are discussed in detail in other texts. Table 7-1 lists normal complete blood count values. Table 7-2 lists normal blood chemistry values for horses.

TABLE 7-1

Normal Complete Blood Count Values

Packed cell volume (%)	32–52 (42)
Hemoglobin (g/dl)	11–19 (15)
Red blood cells (×10⁶ µl)	6.5–12.5 (9.5)
Total protein (g/dl)	6.0–8.0
White blood cells (×10³/µl)	5.5–12.5 (9)
Platelets (×10⁵/µl)	1–6 (3.3)
Mean corpuscular volume (MCV) (fl)	34–58 (46)
Mean corpuscular hemoglobin (MCH) (pg)	15.2–18.6
Mean corpuscular hemoglobin concentration (MCHC) (g/gl)	31–37 (35)
Bone marrow myeloid-to-erythrocyte (M : E) ratio	0.94–3.76 : 1 (1.64 : 1)
DIFFERENTIAL, ABSOLUTE
Segs	30%–65% 2700–6700 (4700)
Bands	0%–2% 0–100 (2)
Lymphocytes	25%–70% 1500–7500 (3500)
Monocytes	0.5%–7% 0–800 (400)
Eosinophils	0%–11% 0–925 (375)
Basophils	0%–3% 0–170 (50)

TABLE 7-2

Normal Blood Chemistry Values

Blood urea nitrogen (mg/dl)	10.4–24.7
Creatinine (mg/dl)	0.9–2.0
Glucose (mg/dl)	62.2–114
Albumin (g/dl)	2.5–3.8
Total bilirubin (mg/dl)	0.3–3
Aspartate aminotransferase (AST) (µ/L)	115.7–287
γ-Glutamyltransferase (GGT) (µ/L)	2.7–22.4
Creatine kinase (µ/L)	34–165.6
Alkaline phosphatase (µ/L)	70.1–226.8
Lactate dehydrogenase (DH) (µ/L)	102.3–340.6
Sodium (mEq/L)	133.3–147.3
Potassium (mEq/L)	2.8–4.7
Chloride (mEq/L)	97.2–110.1
Calcium (mg/dl)	10.4–13.4
Phosphorus (mg/dl)	2.3–5.4
Magnesium (mg/dl)	1.8–2.7

Blood Gas Samples

Blood gas analysis is used most often in assessing patients with respiratory disease and monitoring patients under general anesthesia. The analysis determines the oxygen content, carbon dioxide content, and pH of the blood sample. Arterial blood is usually preferred to venous blood for blood gas analysis because it more accurately reflects the ventilation status of the animal. Blood gas analysis as an anesthetic monitoring tool is more common in large than small animals.

Technician Note

Arterial blood is often preferred to venous blood for blood gas samples because it more accurately reflects the ventilation status of the animal.

Samples for blood gas analysis must not clot and must not be contaminated by atmospheric air. Typically, a 25-ga needle and 3-ml syringe are used to obtain the sample. To prevent coagulation of blood, just enough heparin is aspirated to fill the needle and appear in the needle hub. No air or air bubbles should be in the syringe. At least 1 ml of blood is drawn from an artery or vein, and the needle is immediately capped with a rubber stopper to keep air from contacting the sample (rubber stoppers from blood tubes work well). The syringe/needle/cap combination is placed on ice and promptly analyzed. The sample should be run within 10 minutes but may yield accurate results for about hours if the sample is maintained on ice and kept airtight. The patient’s temperature should be taken at the time of collection, and the analysis must be corrected for body temperature (Fig. 7-2).

FIGURE 7-2 Blood gas collection from the metatarsal artery in a neonate. A small diaper dampened with water and placed in the microwave for 20 seconds was placed over the artery for 30 seconds before collection. The warm compress allowed for dilation and easy visualization of the artery. Topical application of a local anesthetic cream may decrease movement before collection. (From McAuliffe SB, Slovis NM: *Color atlas of diseases and disorders of the foal.* St. Louis, WB Saunders, 2008.)

Technician Note

Samples used for blood gas analysis cannot be exposed to atmospheric air or allowed to clot.

Urine Collection

The purpose of urine collection is to obtain a sample of urine for laboratory analysis. Indications for urine collection include urinary tract disease, various systemic diseases, and toxicologic/pharmacologic analysis. Cystocentesis is not feasible in large animals; therefore, sampling is limited to voided urine collection and bladder catheterization. Cystocentesis is not used in adults because of the inability to stabilize the bladder and the possibility of perforating the intestines during the procedure. Cystocentesis has been successfully performed in foals and small ponies/miniature horses, but it is a risky procedure and not often attempted.

Technician Note

Due to the size of the animal and possible complications, cystocentesis is not often performed in the large animal.

Voided urine is not sterile and therefore is not suitable for culture and sensitivity testing. Urine is collected into a clean container when the horse urinates. When catching a sample of voided urine, it is best to avoid the initial urine stream and collect a midstream sample. The initial urine stream tends to contain more mucus and cell debris and may not be representative of the content of the urine. Urination is sometimes facilitated with diuretic drugs such as furosemide to reduce the time spent waiting for a voided sample. When analyzing diuretic-induced urine samples, the effects of the drug on parameters such as urine specific gravity must be considered. To prepare for a voided collection, you will need a clean container, examination gloves, and possibly diuretic drugs. Some racehorses are trained to urinate on command. Each horse is trained with a different command, so it is a good idea to ask the client about the horse’s training. To begin, the horse should be restrained. If diuretics are to be administered, they should be given now. Once the horse begins to urinate, allow the initial portion of the urine stream to pass, then catch the midstream urine in a clean container; finally, cap the container.

Technician Note

• If a diuretic is used prior to collecting a urine sample, the effects of the diuretic on urine specific gravity should be considered.

• When catching voided urine, it is best to collect a midstream sample.

Catheterization of the bladder risks contamination of the urine with bacteria that accumulate on the catheter as it passes through the urethra. Therefore, culture results of catheterized samples must be interpreted with caution. Because cystocentesis is not feasible in large animals, catheterized urine samples are the accepted compromise for microbial testing. Urine collection using catheterization is a sterile procedure, so you should prepare the following equipment before the procedure: a urinary catheter (stallion catheter for males; mare catheter or metal Chambers catheter for females [Fig. 7-3]), a 60-ml syringe (catheter or Luer tip depending on the catheter used), sterile gloves, sterile lubricating jelly, and a sterile collection container (plastic or glass).

FIGURE 7-3 *Left,* Flexible female urinary catheter and stylet. *Right,* Flexible male urinary catheter and stylet.

Technician Note

Culture results from catheterized samples must be interpreted with caution because bacteria can accumulate on the catheter as it passes through the urethra.

To begin, the horse should be restrained standing, but the procedure can be performed with the animal in lateral recumbency.

Males

Tranquilization is usually required to cause relaxation and extension of the penis. This procedure induces discomfort; therefore, personnel should position themselves cranially to avoid being kicked with the hindlegs. Additional physical restraint may be necessary. Catheterization of the bladder is performed through the urethra. In males, the urethral entrance is on the end of the glans penis. Following extension of the penis, the tip of the glans penis is prepared with at least three applications of antimicrobial soap/water rinses. The tip of the catheter should be well lubricated with a sterile lubricant. The urethral opening is identified, and the lubricated catheter is passed into the urethra and slowly advanced into the bladder. If the catheter has a stylet, it must be withdrawn to allow urine flow through the catheter. Urine sometimes flows freely from the catheter and may be collected in a sterile container. If urine does not flow freely, a sterile 60-ml syringe is used to aspirate urine from the bladder (Fig. 7-4).

FIGURE 7-4 Indwelling urinary catheter in place following surgery to repair a bladder wall defect. The condom fitted at the end of the catheter acts as a one-way valve that allows urine to flow out but prevents air from entering and inflating the bladder. (From McAuliffe SB, Slovis NM: *Color atlas of diseases and disorders of the foal.* St. Louis, WB Saunders, 2008.)

Technician Note

The penis should be prepped with at least three applications of antimicrobial soap/water rinses.

Females

The tail is tied or held to the side, and the perineum is prepared with at least three applications of antimicrobial soap/water rinses. The clinician’s gloves should be well lubricated with a sterile lubricant. Physical restraint may be necessary; sedation is occasionally necessary. In females, the urethral entrance is located in the floor of the vestibule–vagina junction (Fig. 7-5). The procedure from this point is carried out similar to that of male catheterization.

FIGURE 7-5 Placement of a urinary catheter in a filly foal. (From McAuliffe SB, Slovis NM: *Color atlas of diseases and disorders of the foal.* St. Louis, WB Saunders, 2008.)

After the procedure has been performed, all soap residues should be removed to prevent scalding of the skin and mucosal surfaces. Catheterization may cause temporary irritation of the urethra, leading to increased frequency of urination for 1 to 2 days.

Technician Note

It is important to inform clients that urinary catheterization may lead to increased frequency of urination for 1 to 2 days.

Common procedures include the following evaluations and measurements:

• Gross evaluation: Normal horse urine is clear to cloudy and yellow. The cloudiness is due to a relatively large amount of mucus, which is normal in the horse.

• Urine specific gravity

• Chemical analysis: Determined using reagent test strips (“dipsticks”) and/or machine analysis.

• Urine sediment evaluation: Calcium carbonate crystals are normal and common in horse urine.

• Microbial culture and sensitivity: Performed if infectious disease is suspected or confirmed.

Technician Note

Calcium carbonate crystals are normal and common in horse urine.

Fecal Collection

Fecal collection is performed for parasitic evaluation and/or microbial culture. Indications for fecal collection include suspected intestinal parasite infestation or suspected intestinal bacterial/viral/protozoal infection.

Feces can be collected from the ground using a glove or clean container or from the rectum using a glove or rectal sleeve. A hand is inserted into the glove/sleeve and used to grasp the feces. Then, while maintaining a grasp on the feces with the hand, the glove/sleeve is simply turned inside out. This maneuver keeps the feces inside the glove/sleeve, and physical contact with the fecal material is avoided. The glove/sleeve is tied in a knot above the feces and transported to the laboratory. Sample sizes for large and small animals are similar; large animals do not require voluminous fecal samples.

Technician Note

The same amount of fecal material is needed for a large animal fecal analysis as is needed in small animals.

The following observations should be made when performing a fecal laboratory analysis:

• Gross evaluation: Normal equine feces are mostly solid, in formed fecal balls. Color is light to dark green. Mucous coating may indicate that the feces have been retained longer than normal.

• Parasitic evaluation: Fecal flotation ± quantitative techniques ± larval culture.

• Fecal culture/sensitivity: This is especially important when Salmonella is suspected. Salmonella testing usually requires daily samples for 3 to 5 days; at least 10 g of feces should be submitted for each sample.

Abdominocentesis (Abdominal Tap)

The purpose of an abdominocentesis is to obtain a sample of abdominal (peritoneal) fluid for analysis. The fluid is produced by the cells of the peritoneum, which line the abdominal wall and the outer surfaces of the abdominal organs. Abnormalities of abdominal organs may change the character of the abdominal fluid, providing clues for diagnosis of abdominal disease.

The procedure can be readily performed in the clinic or in the field. Indications for abdominocentesis include abdominal disease, gastrointestinal (GI) or non-GI origin, and chronic weight loss.

Technician Note

Some indications for abdominocentesis are abdominal disease, GI or non-GI origin, and chronic weight loss.

It is important that all of the appropriate equipment be prepared before beginning the procedure. You will need sterile gloves, ethylenediamine tetraacetic acid (EDTA) and serum (plain) blood tubes, a needle (18- or 19-ga × inch for most adults, 18-ga × 3-inch spinal needle for large or obese horses, 20-ga × 1 inch for foals), blunt trocar or cannula for severely bloated patients, no. 15 scalpel blade if using a trocar or cannula, local anesthetic/syringe/25-ga × -inch needle for anesthesia of the skin (for foals or if using a trocar or cannula), sterile 4 × 4 gauze if using a trocar or cannula, and clean ground cloth/towel for foals.

Adults are restrained in a standing position for the procedure; foals are restrained in lateral recumbency. The procedure is usually performed at the most dependent (lowest) point of the abdomen, on ventral midline or slightly to the right of ventral midline (Fig. 7-6).

The hair should be clipped and sterile skin prep performed (Fig. 7-7). Performing the procedure without clipping the hair is possible, but special care must be taken to properly prepare the area, and the client should be warned about the slightly increased risk for introducing infection.

FIGURE 7-7 Abdominocentesis being performed in the left ventrolateral abdomen. This site was chosen in this foal based on ultrasound findings of a localized peritonitis. (From McAuliffe SB, Slovis NM: *Color atlas of diseases and disorders of the foal.* St. Louis, WB Saunders, 2008.)

Local anesthesia is seldom necessary in most patients when hypodermic or spinal needles are used. However, local anesthesia of the skin is routinely performed for foals or whenever a trocar or cannula is used. A final skin prep is performed after the anesthetic is injected subcutaneously.

Patient restraint is sometimes necessary, although the procedure is well tolerated and many horses require minimal, if any, restraint. The patient should be prevented from walking during the procedure.

Personnel should stand as far cranially as possible to avoid being kicked during the procedure. Patients typically flinch when the needle is passed through the skin but rarely kick.

A small stab incision through the skin is performed if the trocar or cannula method is being used; syringe needles do not require a skin incision. The clinician advances the needle/trocar/cannula until fluid is obtained. Fluid is collected by gravity flow into the sample collection tubes. The EDTA tube sample should be collected first because most of the laboratory analysis will use this sample. At least 1 ml of fluid should be obtained to prevent false laboratory results, which can occur if the ratio of EDTA to abdominal fluid is too high. The serum (plain) tube is used to collect samples for culture if bacterial disease is suspected; as little as one drop of fluid is sufficient for this use. Abdominal fluid may be difficult to obtain, sometimes requiring several attempts.

Bleeding is common after the needle is removed. This bleeding is due to incidental perforation of skin vessels, which are difficult to avoid because they can rarely be seen. Manual pressure over the site stops the bleeding. If a stab incision was made, the clinician may elect to place a simple skin suture or skin staple to close the incision.

The area should be cleaned gently and antibiotic ointment applied daily for 2 to 3 days. Complications such as infection or abscessation of the site are uncommon.

Common procedures include the following evaluations and measurements:

• Gross visual examination: Normal abdominal fluid is pale yellow, clear, and odorless. Its consistency is similar to water.

• Total protein level

• Red blood cell (RBC) and white blood cell (WBC) counts

• Packed cell volume

• Microscopic evaluation

• Cytology: Direct smear, air dried, stained

• Gram stain if bacteria are observed

• Microbial culture and sensitivity: Performed if infection is suspected or confirmed.

• Ancillary tests: May include pH, enzyme, and other chemistries; seldom necessary.

Arthrocentesis (Joint Tap)

Arthrocentesis is performed to obtain synovial fluid from a synovial joint for analysis. All synovial joints contain synovial fluid, which is produced by the cells of the synovial membrane. Disease of the joint often changes the characteristics of the fluid, and the sample can be used to aid diagnosis of various joint diseases and tendon sheath/bursal disease.

Technician Note

Arthrocentesis can be used to aid in diagnosis of various joint diseases and tendon sheath/bursal disease.

Tendon sheaths and bursae are also synovial structures lined by a synovial membrane that produces synovial fluid. Many tendon sheaths and bursae can be accessed for fluid collection in the same manner as the joints. Analysis of these samples is similar to that of joint fluid.

Arthrocentesis and centesis of other synovial structures can be performed in the clinic or in the field. It is common practice to use the procedure to also medicate the synovial structure. After the fluid sample has been obtained, the selected medication is injected into the structure before the needle is removed.

Technician Note

It is common practice to use arthrocentesis to also medicate the synovial structure. The medication is injected before the needle is removed.

Before beginning the procedure it is important to collect all of the following supplies in order to perform the procedure: sterile gloves, EDTA and serum (plain) blood tubes, a needle (depends on many factors, especially location and depth of the joint, size of the patient, temperament of the patient, and clinician preference), sterile 6- or 12-ml syringe, local anesthetic/syringe/needle (if skin block is necessary, e.g., foals, deep synovial structures, and patients that resist the procedure), and the medication to be injected if applicable.

Technician Note

Local anesthesia can be needed for arthrocentesis of foals, deep synovial structures, or patients that resist the procedure.

Patient restraint depends on many factors, including the anatomical location of the synovial structure, the restraint method, and the clinician’s preference. Restraint is required for most cases and ranges from minimal restraint to more severe physical restraint. In some cases, chemical sedation or even general anesthesia is necessary. Most arthrocentesis is performed with the horse standing and weight bearing, although some structures are accessed with the leg elevated. Some synovial structures are accessible from more than one location (Fig. 7-8). The patient must be motionless while the needle is in the joint because motion can bend or break the needle, and the cartilage and synovial membrane surfaces of the joint can be severely lacerated or punctured.

FIGURE 7-8 Arthrocentesis of the fetlock joint. A, The joint can be approached through the lateral aspect of the palmar or plantar pouch. B, The fetlock joint also can be entered cranially on either side of the extensor tendon. The fetlock is readily entered at this site when there is marked effusion of the joint. The advantage of this site is that inadvertent penetration of the metacarpal or metatarsal blood vessels is less likely. (From McAuliffe SB, Slovis NM: *Color atlas of diseases and disorders of the foal.* St. Louis, WB Saunders, 2008.)

The hair should be clipped and the skin sterilely prepped. It is possible to perform the procedure without clipping the hair, but special care must be taken to properly prepare the area, and the client should be warned about the slightly increased risk for introducing infection. In either case, after the final alcohol scrub, I recommend applying povidone–iodine solution to the area and allowing it to air dry. The solution remains on the patient until the procedure is completed.

Local anesthesia is usually necessary for the skin and subcutaneous tissue if a large needle is to be used, such as for the hip and shoulder joints. Local anesthesia is sometimes required for horses that resist the procedure. It can be provided as a small anesthetic bleb at the skin puncture site or by regional anesthesia of peripheral nerves on the limbs. Most horses do not require local anesthesia for lower leg arthrocentesis.

Technician Note

Local anesthesia is usually necessary for the skin and subcutaneous tissue if a large needle is used for arthrocentesis of the hip and shoulder joints.

Once the needle enters the synovial structure, fluid may flow freely or not at all. Occasionally the clinician uses a sterile syringe to aspirate the synovial fluid. The sample for the EDTA tube should be collected first because most of the laboratory analysis will use this sample. At least 1 ml of fluid should be obtained to prevent false negative and false positive laboratory results, which can occur if the ratio of EDTA to fluid is too high. The serum (plain) tube is used to collect samples for culture if bacterial disease is suspected; as little as one drop of fluid is sufficient for this use.

Technician Note

At least 1 ml of fluid should be obtained during arthrocentesis if the sample is to be added to EDTA in order to prevent false laboratory results.

After the fluid samples are collected, medication can be injected through the needle into the synovial space.

Bleeding from the skin is not unusual after needle withdrawal and is controlled with direct pressure. If bleeding occurs, the blood is cleaned from the skin and antibiotic ointment is placed over the injection site. Bandaging depends on several factors. Often the joint disease itself requires some form of bandaging. Otherwise, many clinicians prefer to cover the injection site for 24 hours to minimize the risk of infection. This practice becomes more essential with structures closer to the ground (i.e., the distal limbs).

Technician Note

Many clinicians prefer to bandage the injection site for 24 hours after arthrocentesis to help minimize the risk of infection.

Exercise is also dictated by the nature of the joint disease, the horse’s occupation, and the type of medication injected into the joint (if this was performed).

Common procedures include the following evaluations and measurements:

• Gross visual examination: Normal synovial fluid is yellow, clear, and odorless. The fluid should have viscosity (i.e., be “stringy”); watery consistency is abnormal.

• Total protein level

• RBC and WBC counts

• Packed cell volume

• Microscopic evaluation

• Cytology: Direct smear, air dried, stained

• Gram stain if bacteria are observed

• Microbial culture and sensitivity: Performed if infection is suspected or confirmed.

• Ancillary tests: May include pH, enzymes, chemical mediators of inflammation, and other chemistries; cartilage fragment analysis and mucin clot tests are no longer routinely performed.

Cerebrospinal Fluid Collection

CSF is collected from the subarachnoid space for analysis. CSF is produced by ependymal cells in the ventricles of the brain and flows in the subarachnoid space around the brain and spinal cord. CSF also flows through the ventricles of the brain and the central canal of the spinal cord but cannot be safely accessed in these locations. Collection of CSF is used to diagnose brain and spinal cord disease and to differentiate peripheral nervous system disease.

Technician Note

CSF collection is used to diagnose brain and spinal cord disease and to differentiate peripheral nervous system disease.

Diseases of the central nervous system may produce changes in the character and composition of CSF; therefore, CSF is used as a diagnostic aid in neurologic disease.

CSF in the subarachnoid space can be readily accessed for sampling at two locations: the atlantooccipital space (also referred to as the cisterna magna) and the lumbosacral space. These procedures are significantly different in technique, but the CSF obtained from either location is essentially the same, and there is no difference in laboratory analysis procedures.

Before beginning the procedure, collect the following equipment: sterile gloves, sterile 12- or 20-ml syringe, EDTA and serum (plain) tubes, a needle (18-ga × 3-inch spinal needle for atlantooccipital tap [Fig. 7-9], 8-inch spinal needle for lumbosacral tap), local anesthetic/12-ml syringe/20-ga × -inch needle for lumbosacral tap or general anesthesia (injectable or inhalation) for atlantooccipital tap.

FIGURE 7-9 An 18-gauge × 3-inch spinal needle.

Atlantooccipital Space Collection

The atlantooccipital space can only be safely accessed with the patient under general anesthesia. Although the procedure carries the risks of general anesthesia, it is a brief procedure, and injectable anesthesia can be used. CSF is technically easy to obtain at this location. If the patient is ataxic, as many neurologic patients are, the risk of self-injury during the anesthetic recovery period is increased and may be unacceptable. Atlantooccipital space is located just caudal to the poll, on the dorsal midline, at the level of the wings of the atlas (Figs. 7-10 and 7-11). This procedure must be performed with the horse under general anesthesia; therefore, proper preparation for anesthesia is required. The patient is placed in lateral recumbency. A rope is usually placed around the nose and pulled caudally to ventroflex the head and neck; this opens up the atlantooccipital space and facilitates needle placement. The patient is clipped and sterilely prepped for the procedure. The needle is advanced by the clinician, paying careful attention to anatomical landmarks, until the bevel is confirmed to be in the subarachnoid space. With atlantooccipital taps, fluid usually flows freely from the needle and can be collected from the needle hub. Fluid may flow freely from the lumbosacral space but usually needs to be collected by gentle aspiration with a sterile syringe because it cannot be easily collected by gravity flow with the needle in its vertical position. Fluid is usually collected into both EDTA and serum tubes; more than 1 ml should be placed in the EDTA tube.

FIGURE 7-10 Cerebrospinal fluid collection from the atlantooccipital space.

Technician Note

• CSF collection from the atlantooccipital space requires general anesthetic risk.

• The atlantooccipital space is located just caudal to the poll, on the dorsal midline, at the level of the wings of the atlas.

Lumbosacral Space Collection

The lumbosacral space is usually accessed in the awake, standing patient; this avoids the risk of recovery from general anesthesia. However, the lumbosacral space is technically more difficult to enter, and patients may display violent reactions to pain from the procedure. Lumbosacral space is located on the dorsal midline, at the level of the wings of the ilium (Fig. 7-12). The procedure is performed in the standing patient, usually with sedation. Care must be taken not to oversedate the horse, which may cause excessive body swaying. Because the patient must stand still during the procedure, stocks are highly desirable to restrict movement. The horse must also stand squarely with weight distributed evenly on all legs because leaning makes the procedure difficult to perform. The patient is clipped and sterilely prepped for the procedure, and local anesthesia of the skin, subcutaneous tissues, and deeper tissues is performed. A final prep is performed after the local anesthetic is injected. The procedure is then performed as if you were collecting fluid from the atlantooccipital space.