Chapter 22
Equine Transfusion Medicine
Margaret C. Mudge1 and Olivia H. Williams2
1The Ohio State University, Department of Veterinary Clinical Sciences, Columbus, Ohio, USA
2Piedmont Equine Associates, Madison, Georgia, USA
Introduction
Transfusions of whole blood (WB) or packed red blood cells (PRBCs) are indicated for the treatment of acute hemorrhage or severe anemia in horses. While blood transfusions are not a frequent procedure performed in horses, practitioners should be prepared to collect and administer blood as a life-saving measure. There are some important differences in the collection, pre-transfusion testing, and administration of blood to horses compared to small animals. This chapter will provide information to help with deciding when to transfuse, as well as the practical steps for the collection and administration of blood to horses.
Indications for red blood cell transfusions
WB transfusions provide additional oxygen-carrying capacity, coagulation factors, proteins, and blood volume. WB transfusions are indicated for horses that have suffered acute blood loss from trauma, surgery, or other conditions such as splenic rupture or uterine artery hemorrhage. In horses that have normal blood volume with severe anemia, PRBCs would be the most appropriate product, although WB transfusions are often given when blood product processing is not available or clotting factors are also needed. Plasma transfusions are commonly administered to foals to supplement low immunoglobulin levels or to horses with coagulation disorders.
Transfusion triggers
Specific physical examination and clinicopathologic values that indicate the need for blood transfusion are often termed “transfusion triggers”. While there are no set variables that serve as transfusion triggers in horses, a combination of physical examination and clinicopathologic parameters (Box 22.1) can be used to guide the decision to transfuse (Hurcombe et al. 2007).
Physical examination and estimation of blood loss
Physical examination findings such as pale mucous membranes, tachycardia, tachypnea, sweating, colic, and lethargy can indicate need for a blood transfusion, especially when blood loss is estimated to be greater than 30% of blood volume. Acute blood loss can result in hypovolemic shock in addition to loss of red cell mass, so findings might also include cold extremities, hypotension, and increased blood lactate concentrations. In the early stages of hemorrhage, packed cell volume (PCV) can still be normal due to a lack of fluid redistribution or replacement, as well as splenic contraction. The PCV and total protein (TP) will decrease as fluid redistributes from the interstitial to the intravascular space. Some fluid redistribution occurs almost immediately after blood loss, although it might take hours to days for complete restoration of blood volume (Drucker et al. 1981; Ryan et al. 2012; Saito et al. 2013). While decreased PCV is a useful indicator of blood loss in most patients, it tends to underestimate acute hemorrhage.
In a group of awake horses with mild blood loss, heart rate and blood lactate increased, while central venous pressure decreased (Magdesian et al. 2006). In a study of anesthetized horses with massive hemorrhage, PCV actually increased by the end of the period of blood loss, presumably due to a sympathetic response and release of red blood cells (RBCs) from the spleen. The horses also had stable heart rates, making this parameter an unreliable measure of blood loss in anesthetized horses. Pale mucous membranes with prolonged capillary refill time, decreased TP, hypotension, and hypoxemia are better indicators of blood loss in anesthetized horses (Wilson et al. 2003).
Estimating blood loss is important during the early stages of hemorrhage when PCV cannot be used as a reliable guide. Table 22.1 provides guidelines for estimating acute blood loss in horses. A blood transfusion is indicated if physical examination parameters indicate blood loss approaching 30% of total blood volume. If intravenous (IV) fluids are given for resuscitation, the PCV and TP will decrease more rapidly. A blood transfusion is likely needed if the PCV decreases below 20–25% during an acute bleeding episode, although in acute severe cases a transfusion might be needed before there is a significant decrease in PCV.
Table 22.1 Parameters associated with estimated blood loss in adult horses (American College of Surgeons 2004)
| Blood loss (% blood volume) | Heart rate (beats/min) | Respiratory rate (breaths/min) | Capillary refill time | Blood pressure | Urine output | Other physical examination findings |
| Up to 15% | 32–48 | 8–16 | 1–2 seconds | Normal | Normal | Possible mild anxiety |
| 15–30% | 50–60 | 20–30 | 2–3 seconds | Normal | Mildly decreased | Mild anxiety |
| 30–40% | 60–80 | 30–40 | 3–4 seconds | Decreased | Decreased | Altered mentation; cool extremities |
| >40% | >80 | >40 | Absent, very pale mucous membranes | Severe hypotension | Negligible | Obtunded; cool extremities |
Oxygen extraction
Oxygenation status can also help to determine the need for blood transfusions during acute hemorrhage or chronic anemia. An increase in blood lactate concentration despite volume replacement with crystalloids or colloids can indicate continued tissue hypoxia and the need for a blood transfusion (Greenburg 1995; Magdesian et al. 2006). Oxygen extraction ratios are also useful measurements; a ratio greater than 40% in the context of blood loss might indicate a need for a blood transfusion (Magdesian 2008).
PCV and TP
In equine patients with chronic hemolytic anemias, PCV and TP can be useful indicators of the need for a blood transfusion. While there is no set transfusion trigger for horses, a PCV less than 12–15%, especially in conjunction with the previously mentioned physical examination findings (e.g., pale mucous membranes, tachycardia, tachypnea, lethargy), represents an indication for a blood transfusion. Transfusions might be required for horses with a higher PCV if concurrent disease such as a respiratory condition or sepsis is present. Since animals with chronic anemia are normovolemic or even hypervolemic, PRBCs are preferred to lessen the volume administered, although WB can also be used if PRBCs are not available.
For acute and chronic anemias, the primary goal of the blood transfusion is to increase the oxygen-carrying capacity. RBCs from allogenic transfusions have a much shorter half-life compared to autologous red cells, so transfusion should still be considered a temporary measure to restore oxygen-carrying capacity. Full resolution of the anemia will require the horse’s erythropoietic response and successful treatment of the underlying disease.
Indications for plasma product transfusions
Plasma products are commonly administered to neonatal foals to treat failure of transfer of passive immunity, otherwise termed failure of passive transfer (FPT). Plasma transfusions are also used for the treatment of clotting factor deficiencies, hypoalbuminemia, and disseminated intravascular coagulation in horses (Welch et al. 1992). Plasma can be considered fresh frozen plasma (FFP) when frozen within 8 hours of collection. FFP contains immunoglobulins, coagulation factors (fibrinogen and factors II, VII, IX, X, XI, and XII), and cofactors (factors V and VIII), as well as the anticoagulant proteins antithrombin, protein C, and protein S. Frozen plasma is plasma that is frozen more than 8 hours after collection and has decreased activity of factors V and VIII. There are a number of commercially available equine plasma products (Table 22.2).
Table 22.2 Commercial sources of equine plasma products
| Company | Products | Website |
| Veterinary Immunogenics (Cumbria, UK) | Hypermune plasma R. equi antibody | www.veterinaryimmunogenics.com |
| Lake Immunogenics (Ontario, NY) | HiGamm R. equi antibody Plasmune Custom antigen select plasma | www.lakeimmunogenics.com |
| Mg Biologics (Ames, IA) | Hyperimmune plasma Normal equine plasma R. equi antibody Designer plasma | www.mgbiologics.com |
| Plasvacc USA (Templeton, CA) | EquiPlas EquiPlasPlus (high IgG) R. equi antibody Custom antibody Antitoxin plasma Whole blood | www.plasvaccusa.com |
Plasma for colloid support
Plasma can be used for colloid support, for example when the TP concentration is less than 
, the serum albumin concentration is less than 
, or the colloid osmotic (oncotic) pressure is less than 14 mmHg. If clotting factors and albumin are not needed, synthetic colloids such as hydroxyethyl starch can be used for oncotic support. Synthetic colloids have been associated with coagulopathies and acute kidney injury in human patients (US Food and Drug Administration 2013), but extensive information regarding these side effects in equine patients is not currently available. Synthetic colloids have been evaluated in horses with colic and colitis, and side effects were not recognized (Bellezzo et al. 2014; Dugdale et al. 2015). However, a study investigating the hemostatic effects of synthetic colloids in healthy mares indicates that administration of tetrastarch (130/0.4) at 40 mL/kg produces thromboelastography changes trending towards hypocoagulation, though the values remained within reference ranges (Viljoen et al. 2014). FFP has not been evaluated specifically for the treatment of hypoalbuminemia in horses.
Plasma for failure of passive transfer of immunity
FPT in neonatal foals older than 12 hours is best treated with plasma transfusion, as colostrum absorption is greatly diminished by this time (Jeffcott 1975). An immunoglobulin G (IgG) concentration <200 mg/dL (2 g/L) is considered complete FPT and an IgG concentration between 400 and 800 mg/dL (4 and 8 g/L) is considered partial FPT. Although plasma transfusions are not always needed for foals with partial FPT, they are recommended for foals that have pre-existing infection or exposure to pathogens such as Clostridium spp.
Commercially available fresh frozen hyperimmune equine plasma is most commonly used for the treatment of FPT in neonatal foals. Hyperimmune plasma is made by performing plasmapheresis using donor horses that have been hyperimmunized. Equine FFP is a United States Food and Drug Administration (USFDA) licensed product; most have a minimum guaranteed IgG concentration (e.g., 
or 
) and a 2–3 year shelf life when frozen. Although commercially available hyperimmune plasma has very high IgG concentrations, plasma from local donor horses might provide better protection against specific local pathogens.
Plasma for specific antibodies
Other available equine plasma products include those with bacterial or viral specific antibodies. Escherichia coli (J5) and Salmonella typhimurium hyperimmune plasma have been evaluated for the treatment of equine endotoxemia, with mixed results (Spier et al. 1989; Durando et al. 1994; Southwood 2004; Peek et al. 2006). The use of Rhodococcus equi hyperimmune plasma for the prevention of R. equi has also been controversial (Madigan et al. 1991; Hurley and Begg 1995; Giguère et al. 2002; Perkins et al. 2002; Caston et al. 2006). Other plasma products available for treatment of specific diseases include botulism antitoxin, West Nile virus antibody, and Streptococcus equi antibody.
Equine blood groups
There are eight recognized equine blood groups, A, C, D, K, P, Q, U, and T, as well as 34 different factors identified within the first seven of these groups (International Society for Animal Blood Group Research 1987). RBC antigens Aa and Qa are most often associated with neonatal isoerythrolysis (NI), and Ca antibodies are commonly found in horses that lack the Ca antigen. Horses of the same breed are more likely to have similar blood types, but many breeds display a diverse array of polymorphisms (Bowling and Clark 1985). Blood typing used to be an accepted method of determining parentage in horses, but this method has been replaced by DNA typing.
Compatibility testing
In an emergency situation, a blood transfusion can be given for the first time with very minor risks of serious transfusion reactions since horses rarely have naturally occurring antibodies. Horses that are negative for Aa and Qa and lack RBC antibodies are preferred blood donors when a crossmatch cannot be performed. If typed donor horses are not available, a healthy gelding of the same breed should be used. Horses can develop alloantibodies within 1 week of transfusion, so blood typing and crossmatching are recommended before a second transfusion is performed (Wong et al. 1986). However, a second blood transfusion can be performed safely within 2–3 days of the first transfusion without a crossmatch. In non-emergency situations (or when testing is readily available), blood typing and crossmatching are recommended to limit the risk of immunologic transfusion reactions and increase the chance of optimal survival of transfused RBCs.
Neonatal isoerythrolysis
Blood compatibility testing is often used to prevent or diagnose NI. The mare is usually sensitized to a different blood type during pregnancy (exposure to the stallion’s blood type that is shared by the fetus) or occasionally during a blood transfusion. The anti-RBC antibodies are transferred to the foal when it ingests colostrum. When there are sufficient RBC antibodies, the foal’s RBCs are lysed. The majority of NI cases have been attributed to the Aa and Qa antigens, with Qrs, Qb, Qc, Da, Db, Dc, Ka, Pa, and Ua also reported to cause NI (MacLeay 2001).
Blood typing (see below) of the mare and stallion can be used to determine whether there is any risk of NI. While it is common for the mare and stallion to have different blood types, the likelihood of NI is quite low (less than 2%). A more specific test for risk of NI would be to measure RBC antibodies in the mare 1–2 weeks prior to foaling. After the foal is born, a crossmatch of the foal’s blood and mare’s serum (minor crossmatch to detect anti-foal RBC antibodies in the mare’s serum) or the foal’s blood and mare’s colostrum (the so-called “jaundiced foal agglutination test”) can be performed.
Blood typing
Rapid blood typing test kits are not commercially available, so blood must be sent to a specialized testing laboratory (Box 22.2). Since blood typing is not readily available and often not possible for emergency situations, performing blood typing and measuring alloantibodies is usually only done when screening equine blood donors or testing for risk of NI. Blood typing involves the use of antisera to detect specific RBC antigens. Blood typing and antibody screening prior to initial transfusion are more important for horses when subsequent blood transfusions are anticipated or for broodmares that can produce foals with NI if sensitized to other blood group factors (Wong et al. 1986).
In practice, transfusion recipients are not usually blood typed. Instead, blood donors are blood typed and ideally are Aa and Qa negative. A rapid agglutination method for detection of equine RBC antigens Ca and Aa has been developed (Owens et al. 2008). If this test becomes commercially available, it would be more practical than complete blood typing for pre-transfusion testing.
Antibody screening
Ideally, donor horses should be screened for alloantibodies yearly. Naturally occurring anti-Aa and anti-Ac antibodies can be measured in horses and are usually agglutinin antibodies. Mares that have been previously sensitized during pregnancy can have anti-Aa hemolysins as well. Recipients can also be tested for RBC antibodies, but such testing is not widely available and is usually impractical in the emergency setting. Antibody screens are routinely performed in mares that are at risk of having anti-RBC antibodies to the foal’s blood type and are more likely to cause NI.
Crossmatch
A crossmatch is recommended prior to blood transfusions, especially for horses that might have been exposed to RBC antigens during a previous transfusion or parturition. Hemagglutination crossmatching is widely available and rapidly performed, but it will not predict all immunologic transfusion reactions, namely hemolytic reactions. The major crossmatch detects agglutination between the donor’s RBCs and the recipient’s plasma. The minor crossmatch detects agglutination between the donor’s plasma and the recipient’s RBCs.
Equine crossmatches can be difficult to interpret due to rouleaux formation. Normal rouleaux should disperse when a small amount of saline is mixed with the blood, whereas agglutination will not disperse (Figure 22.1). Crossmatching can also be difficult to interpret in horses with immune-mediated hemolytic anemia and autoagglutination. The procedure for performing a hemagglutination crossmatch is described in Box 22.3. Although stored blood is convenient to use for the crossmatch, fresh blood samples yield more reproducible results. Even with sample storage of only 1 week, crossmatches have poor reproducibility compared to freshly drawn blood samples, which could potentially lead to the exclusion of compatible donors (Harris et al. 2012).
Figure 22.1 Normal rouleaux formation of equine red blood cells (a) is differentiated from Agglutination seen with an incompatible crossmatch (b).
The routine crossmatch evaluates agglutination, but does not test for hemolytic reactions. Rabbit complement can be added to the reaction mixture for hemolytic testing, but this is not routinely performed as part of the crossmatch except in some laboratories (Becht et al. 1983). Crossmatch testing also does not accurately predict the lifespan of the transfused RBCs or the development of antibodies to the transfused RBCs. As such, transfusion reactions have been reported even with a compatible crossmatch (Hurcombe et al. 2007). If the minor crossmatch is incompatible, but the major crossmatch is compatible, the transfusion can still be performed after washing the donor RBCs and transfusing PRBCs only.
Donor selection
An ideal equine blood donor is a healthy young adult (3–16 years old) gelding with a good temperament. The donor should also have a lean body weight of at least 450 kg, a minimum PCV of 35%, and a total protein 6.0 g/dL (60 g/L). Geldings are preferred over mares, since mares might have RBC antibodies if they were sensitized during gestation. Potential donors with a history of receiving blood products should be excluded during the selection process, as this increases the likelihood of having circulating alloantibodies. Horses selected as donors should also test negative for equine infectious anemia and have an up-to-date vaccination status that includes vaccinations against rabies, tetanus, rhinopneumonitis, Eastern and western encephalitis, and West Nile virus. Donors that are used for production of USFDA-licensed plasma products must also be tested for piroplasmosis, dourine, glanders, and brucellosis (American Association of Equine Practitioners 2009).
Donor blood groups
Because of the large number of blood groups and factors, there are no true universal equine donors. The RBC antigens Aa and Qa are the most immunogenic and have been commonly associated with NI, so the ideal donor should lack the Aa and Qa antigens. There are breed-specific blood factor frequencies, so a donor of the same breed as the recipient is preferable, especially when blood typing is not available. Donkeys have a RBC antigen known as “donkey factor”, which is not present in horses. Therefore, donkeys or mules should not be used as donors for horses, as the transfused horses can develop anti-donkey factor antibodies (McClure et al. 1994). Horses without anti-donkey factor antibodies can be used as donors for donkeys and mules. For foals with NI, the mare can be used as a blood donor, but the RBCs must be washed prior to transfusion. An alternative to fresh blood from donor horses is commercially available WB or PRBCs (Table 22.2).
Autologous blood transfusions
When a surgical procedure is planned in advance and there is a high risk of substantial blood loss, preoperative autologous donation should be considered, as the horse is its own ideal blood donor (Mudge 2005). The half-life of transfused autologous RBCs after 28 days of storage is approximately 30 days, compared to a 20-day half-life for fresh, crossmatched, blood typed, allogeneic blood (Owens et al. 2010; Mudge et al. 2012). Intraoperative or post-hemorrhage cell salvage is also an option for autologous transfusion, and its use has been reported in a horse with post-castration hemorrhage (Waguespack et al. 2001). In these situations, RBC recovery can be performed with specialized cell salvage equipment, which washes and filters collected blood (see Chapter 21). Cell salvage can also be performed with anticoagulation and simple filtration (Waters 2005). The technique of cell salvage is limited to cases when the salvaged blood is not in an area of infection or malignancy, unless specialized washing and filtering equipment is used. Automated cell salvage has not been reported in horses, but has been reported in small animal patients (Kellett-Gregory et al. 2013).
Blood collection technique
When a transfusion is anticipated and fresh WB will be used, the donor horse should be weighed and the PCV and TP measured prior to blood collection (Figure 22.2). Ideally, the donor PCV should be >35%. The maximum volume collected is 20% of the horse’s blood volume (approximately 16 mL/kg body weight), which should be calculated based on lean body weight (Malikides et al. 2001). Blood is collected from the jugular vein of the donor horse using either direct needle cannulation or catheterization. When a large volume of blood is needed, a 10- or 12-gauge catheter is recommended, although a 14-gauge catheter is also sufficient. Blood flow can be improved by placing the catheter opposite to the venous blood flow (i.e., catheter directed toward the head) (Figure 22.3). Both jugular veins can be used if a large volume of blood is needed immediately. When ≥15% of total blood volume is collected, volume replacement with IV isotonic crystalloid fluids is recommended. The donor horse’s heart rate, respiratory rate, and mentation should be monitored during the blood collection. Vital parameters are expected to normalize within 1 hour of collection.
Figure 22.2 A donor horse is restrained in stocks and has food available to help him remain quiet during the collection process.
Figure 22.3 The intravenous catheter is directed rostrally so that the collection is in the direction of venous blood flow.
Vacuum canisters can be used to speed the collection, but glass bottles with vacuum are not recommended as the glass inactivates platelets and can damage the RBCs (Sasakawa and Tokunaga 1976; Mudge et al. 2004). Commercially available 450 mL blood collection bags can be used in horses (Baxter Fenwal, Deerfield, IL; MWI Veterinary Supply, Meridian, ID). There are also commercially available whole blood collection kits that make the procedure relatively easy (Dynavet Blood/Plasma Collection Kits, set.09, Plasvacc USA, Inc., Templeton, CA). These kits include a collection bag, the appropriate amount of citrate anticoagulant required, and a large bore needle attached to the collection line, which allows for rapid collection via direct jugular venipuncture. A filtered blood administration set for delivery to the recipient is also included. Using the provided needle makes collection simple, but can be more invasive, especially if collecting more than one 2 L bag, as jugular venipuncture will need to occur multiple times. The donor’s jugular veins should be treated with care as they will be used repeatedly over several years, with collections occurring up to every 30 days.
Alternatively, an IV catheter can be placed in the jugular vein and a solution set used as a collection line (Figure 22.4). This allows for multiple bags to be collected without causing repeated trauma to the jugular vein. There are larger (4 L) collection kits available with 12-gauge collection needles Jørgen Kruuse, Havretoften 4, DK-5550 Langeskov, Denmark). Blood collection bags can also be made by the addition of anticoagulant to an empty sterile collection bag. The injection of anticoagulant has the potential to introduce bacterial contamination, so this practice should be avoided, especially when blood storage is planned. The supplies required for blood collection are listed in Box 22.4 and the procedure is outlined in Box 22.5.
Figure 22.4 Dynavet kit with 10 gtt solution set attached for collection using an intravenous catheter.
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