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Antimicrobials are commonly used to treat equine infections, but complications with therapy are also relatively common. Antimicrobial‐associated diarrhoea (Section 6), immune‐mediated diseases and nephrotoxicity (Table ) have been reported as sequelae of antimicrobial therapy in equine practice. This does not indicate that clinicians should avoid using these drugs, only that knowledge of the potential for adverse effects is critical when deciding on the most appropriate antimicrobial to use and can assist in avoiding serious side effects, wherever possible, or at least in detecting and responding to them early. The growing problem of antimicrobial resistance also affects equine veterinarians with increasing frequency. Antimicrobial stewardship and responsible prescribing are essential for a future in which effective antimicrobials are available, as it is unlikely that new antimicrobials will become available for use in horses. Australia's conservative regulatory approach has prevented the registration of antimicrobials (such as 4th generation cephalosporins) that are available elsewhere, and the antimicrobial development pipeline is struggling to attract investment, so few new antimicrobial agents, let alone novel classes, are currently in development.

Resistant pathogens of concern in horses include extended‐spectrum β‐lactamase (ESBL) producing Enterobacteriaceae , methicillin‐resistant Staphylococcus aureus (MRSA) and multi‐resistant Salmonella species, while pathogens with high levels of intrinsic and acquired resistance are also common causes of disease, including Pseudomonas species, Klebsiella species, Enterococcus species and Acinetobacter species. The limited antimicrobial options available for these pathogens make the treatment of cases associated with them extremely challenging. In Australia, antimicrobial importance is assigned by the Australian Strategic and Technical Advisory Group on Antimicrobial Resistance (ASTAG). Antimicrobials are categorised as low‐, medium‐ or high‐importance based on the consequence to human health if resistance to them were to develop in human pathogens and the number of antimicrobial treatment options available. It is essential that high‐importance antimicrobials, such as ceftiofur, enrofloxacin, rifampicin, amikacin and polymyxin B, are used prudently in horses and use should be based on culture and antimicrobial susceptibility testing. Some veterinary boards (e.g. Victoria) require veterinarians to have antimicrobial stewardship protocols in place.

These guidelines comprehensively outline how numerous diseases in horses are linked to bacterial infection. However, systemic inflammation caused by viral or fungal infections, bacterial toxins, tissue damage or neoplasia can mimic the clinical signs of bacterial infection. Interpreting haematological findings and fibrinogen and serum amyloid A (SAA) levels can pose challenges, as they may change in response to both infectious and non‐infectious inflammatory conditions. Leukopaenia is frequently observed in association with inflammation, often originating from the gastrointestinal tract, yet it does not necessarily signify the presence of infection or the need for antimicrobial therapy. Similarly, leukocytosis can occur in inflammatory, infectious and neoplastic conditions or because of corticosteroid administration, and thus has limited diagnostic value. Complementary examinations can be performed to strengthen the suspicion of a bacterial infection, and, if possible, a culture of an appropriate sample or a polymerase chain reaction (PCR) assay can confirm infection. However, it often takes time to perform these tests and obtain results. Given the urgency of treatment in many scenarios, clinicians must often rely on professional judgement to determine the likelihood of bacterial infection.

The traditional concept that ‘a course of antibiotics needs to be finished to avoid development of resistance’ is obsolete and not supported by evidence. In fact, the opposite is true – longer courses of antimicrobial therapy may have a greater impact on development of resistance than shorter courses. In most cases, a short duration of antimicrobial therapy, ranging from a single dose (e.g. pre‐operatively) to a course from 24 to 72 h, is sufficient, with long‐term therapy rarely required. Studies from human medicine have shown marked reductions can be made in the duration of therapy of bacterial infections without compromising clinical cure rates. Despite this growing evidence, limiting the duration of antimicrobial therapy in proven infections can be challenging for clinicians. Fear of treatment failure often leads to prolonged use of antimicrobials, even in patients that appear to have recovered clinically. In the past, return to normal haematological parameters or normal concentrations of acute phase proteins have been used as markers for safe discontinuation of antimicrobial treatment. ‘Treatment to clinical cure’ is the new mantra – return to normothermia, improved appetite, return of normal demeanour and a declining trend in inflammatory markers may be better indicators that further treatment is not necessary. More evidence is urgently needed.

It has also been common practice to initially treat systemically ill horses with injectable antimicrobial drugs for 48–72 h, followed by oral treatment. In most cases, this is not necessary. An alternative approach is stopping antimicrobial treatment after 48–72 h and monitoring the patient closely for another 24 h. Should signs of infection reoccur, such as recurrence of a fever, a decrease in appetite or change in demeanour, treatment can be easily reinitiated – but this is rarely necessary because recurrence is rare. Many non‐antimicrobial therapies can be used effectively to combat and control infection. Debridement, lavage, local drainage and cleaning and use of non‐antimicrobial substances that have antimicrobial properties (e.g. honey) can be more effective than systemic antimicrobial therapy and, in many instances, also combat the formation of biofilms. Many of the recommendations in these guidelines represent extra‐label or off‐label use. Veterinarians should be aware of the regulations in their state or territory. Many of the recommendations in this guide represent off‐label use of antimicrobials. Compliance with the legal requirements of your jurisdiction is your responsibility.

Antimicrobial drugs should not be used as an alternative to atraumatic and aseptic surgical techniques. Intravenous prophylactic antimicrobial administration should be given within 60 min of the first incision. Shorter durations of antimicrobial prophylaxis will drive less antimicrobial drug resistance. There are a plethora of data from human surgery and emerging data from veterinary surgery supporting the use of shorter duration surgical antimicrobial prophylaxis. Antimicrobial drug use drives antimicrobial drug resistance, and this is a problem in animal and human health. Inappropriate use, especially prolonged surgical antimicrobial prophylaxis, is detrimental to the patient and increases the costs associated with surgery without yielding any reduction in rates of surgical site infections (SSIs). Antimicrobials disrupt the intestinal microbiome, which is critical to immune function and tissue healing. Antimicrobial use has also been associated with acute diarrhoea in horses, and this can have a high fatality rate. There are three components of surgical antimicrobial prophylaxis. They are: Drug administration time relative to surgery.

Antimicrobial drugs should never be used to replace meticulous atraumatic and aseptic surgical techniques. Clean surgical procedures should not require prophylaxis, and clean contaminated and contaminated procedures should only require perioperative prophylaxis. Dirty and infected procedures require therapeutic, not prophylactic, antimicrobial therapy (Table ).

Prophylactic antimicrobial administration should be timed so that serum and tissue concentrations exceed the minimal inhibitory concentration (MIC) for the organisms likely to be encountered for the duration of the surgery. Antimicrobials cause maximum suppression of infection if given before bacteria gain access to the tissue. Antimicrobials given at a time that does not result in sufficient tissue concentrations during surgery, such as post‐operatively, do not reduce the risk of surgical site infection. Specific recommendations depend on the agent and route of administration. Intravenous antimicrobials: Administer 30–60 min before surgery. Intramuscular procaine penicillin: 3.5 h before surgery. T max for individual drugs given by different routes can be used to assess optimal timing to achieve peak blood concentration at the time of first incision. If repeat dosing is required for prolonged procedures, the dosing interval can be calculated as twice the elimination half‐life of the antimicrobial. Oxytetracycline and procaine penicillin: redosing not required due to long elimination half‐life.

Small reductions in arterial blood pressure have been documented following intravenous sodium penicillin in healthy horses,, so care should be taken when administering benzyl penicillin to unwell horses, particularly those with endotoxaemia and dehydration or other conditions that cause or predispose to hypotension.

Assess patient signalment and risk factors. Is surgical prophylaxis indicated (Table ). If so, identify the appropriate agent, dose, route and timing of administration to reach MIC for target tissues. Ensure procedural planning so that the agent is given at the appropriate time (IV within 30–60 min of surgery, procaine penicillin IM 3.5 h before surgery). Cease prophylactic medication as soon as possible after surgery (Table ). Other factors to consider that reduce the risk of intraoperative infection include clipping the patient's hair <4 h before surgery and minimising the number of people in the surgical theatre.

Benzyl penicillin at 22,000 IU/kg (12 mg/kg) IV and gentamicin at 6.6 mg/kg IV within 60 min of first incision Procaine penicillin G at 22,000 IU/kg (22 mg/kg) IM 3.5 h before surgery and gentamicin at 6.6 mg/kg IV within 60 min of the first incision

Fever and heart murmur are usually present. Blood culture is important for diagnosis and to direct antimicrobial therapy. Echocardiography is important to identify valve thickening, heart chamber size and degree of valvular regurgitation. Bacterial endocarditis is an uncommon, life‐threatening cardiac disorder in horses characterised by bacterial invasion of the endothelial surface of the valves or the wall of the heart, resulting in fibrinous clots or vegetations that impede normal cardiac function. Although bacterial endocarditis can affect horses of all ages, it is predominantly found in animals <3 years of age. Cases generally present for evaluation of fever of unknown origin. Clinical signs usually consist of fever with an audible heart murmur (Grade 3/6 or louder) on either side of the thorax, with or without tachypnoea, tachycardia, cardiac arrhythmia or signs of congestive heart failure. Vegetative lesions are most commonly found on the mitral valve, followed by the aortic valve. Lesions on the tricuspid valve are less common and pulmonary valve lesions are rare.

Common bacterial isolates are Streptococcus spp., Actinobacillus spp. and Staphylococcus spp.,, but Serratia marcescens , Pseudomonas spp. and Escherichia coli have also been implicated. Cardiac arrhythmias may result from direct extension of the inflammatory lesion into the myocardium or may be secondary to thromboembolic myocardial ischaemia.

Echocardiography can be used to detect thickening of the heart valves. Blood culture is indicated to identify bacteria present and direct antimicrobial therapy. Multiple blood cultures (5 collected at least 12 h apart) may improve the likelihood of detecting a pathogen. Delaying therapy to allow adequate sample collection is recommended because the identity of the pathogens involved is unpredictable, as is their antibacterial susceptibility, and long durations of therapy are required. Haematological analysis reveals leukocytosis, with a mature neutrophilia and elevated fibrinogen and serum amyloid A (SAA). An electrocardiogram is required if an arrhythmia is detected by auscultation, but this is rare. Cardiac troponin I has been established as a sensitive and specific marker of myocardial injury. Although this is not always present, it could be useful for diagnosis and to monitor the response to treatment.

Broad spectrum antimicrobial therapy is indicated until culture and susceptibility results are available. Resolution of clinical signs, decrease in the size of the lesion on echocardiographic examination, and a reduction in leukocytosis, hyperfibrinogenaemia and SAA should determine the duration of therapy, which is likely to be several weeks.

Benzyl penicillin (22,000 IU/kg (12 mg/kg) IV q6h) and gentamicin (6.6 mg/kg IV q24h) Procaine penicillin G (22,000 IU/kg IM q12h) and gentamicin (6.6 mg/kg IV q24h) [Correction added on 18 September 2025, after the first online publication: The frequency for gentamicin was incorrectly stated as ‘q12h’ and has been corrected to ‘q24h’.]

Guarded to poor. Sterilisation of lesions can be difficult to achieve. Marked chamber enlargement, severe deformation of the valve leaflets, severe valvular regurgitation and signs of congestive heart failure are grave prognostic indicators.

Uncommon disease but found secondary to pleuropneumonia. Echocardiography is important for diagnosis. Drainage of pericardial effusion under ultrasonographic guidance. Pericarditis is uncommon in horses and can be secondary to pleuropneumonia or haematogenous spread of bacteria. The pathogens involved reflect these primary diseases. The clinical presentation includes fever, tachycardia and jugular distension. Heart sounds can be heard over a wider than normal area of the thorax, but sounds can be muffled or pericardial friction rubs can be heard. Pericarditis was found more frequently following exposure to Eastern tent caterpillars in the United States but has not been associated with exposure to the processionary caterpillars present in Australia.

Echocardiography is required to visualise fluid in the pericardial sac. Thickening of the pericardium with fibrin, coating the visceral and parietal pericardial surfaces, can be evident. An electrocardiogram (ECG) might reveal electrical alternans, where a regularly repeating change in P, QRS and T morphology is present due to regular movement of the heart in the pericardial fluid. Hematology reveals an inflammatory leukogram with elevated fibrinogen and serum amyloid A (SAA) concentrations. Sample collection is often possible when there is moderate to severe accumulation of fluid. Fluid analysis reveals high total protein concentrations and elevated white cell counts with a predominance of neutrophils. Culture and susceptibility testing should be pursued in every case, as the identity of the pathogen is unpredictable and may not be consistent with pathogens cultured from the pleural space or tracheal washes.

Drainage of the pericardial fluid under ultrasonographic guidance is critical in cases where fluid accumulation is moderate or severe. Collection of fluid for bacterial culture and susceptibility testing and cytological examination should be pursued whenever possible. Broad spectrum antimicrobial therapy is indicated until culture and susceptibility results are available and can be guided by cytological findings. Non‐steroidal anti‐inflammatory drugs are an important component of medical treatment of pericarditis (flunixin meglumine 1.1 mg/kg IV q24h or 0.5 mg/kg IV q12h). Monitoring of pericardial thickening and cardiac chamber dimensions is advisable as pericardial constriction appears to be an important fatal sequela of equine pericarditis that may develop insidiously, one or possibly up to 2 years after diagnosis. There are no published studies on which to base the duration of therapy, but it is typically 2–3 weeks, or until resolution of the pericardial effusion.

Benzyl penicillin (22,000 IU/kg (12 mg/kg) IV q6h) and gentamicin (6.6 mg/kg IV q24h) until culture and susceptibility results are available OR procaine penicillin G (22,000 IU/kg IM q12h) and gentamicin (6.6 mg/kg IV q24h) until culture and susceptibility results are available Therapy is generally administered for 2–3 weeks or until concomitant pleuropneumonia has resolved

Sprayberry and Slovis followed a small group of young thoroughbred horses and found a reasonably good prognosis, with some attaining a high level of athletic performance, after successful treatment of pericarditis. This study was dominated by cases associated with Eastern tent caterpillars, which are not present in Australia, so the prognosis may vary with other primary causes of disease.

Severe disease that often requires intensive care. Aseptic, non‐infectious and not contagious. Purpura haemorrhagica (PH) is a non‐septic, immune‐mediated vasculitis that occurs as a rare complication of strangles (or vaccination against it), due to a type 3 hypersensitivity reaction. Typically, disease develops 2–4 weeks after infection with Streptococcus equi , although it may occur more quickly following vaccination (within a week). Although rare, vaccination with agents containing the SeM protein or avirulent S. equi is associated with an increased risk of PH. Occasionally, PH may occur secondary to infection with another infectious organism. It appears to be caused by the deposition of immune complexes on blood vessel walls following antigenic stimulation. Painful pitting oedema results from a necrotising vasculitis. The head, limbs and trunk are most frequently affected, with petechiation and ecchymoses of the mucous membranes. Severe oedema can result in exudation from the skin surfaces, and sloughing of the skin may occur. The vasculitis can affect other sites, including the gastrointestinal tract (resulting in colic), lungs (resulting in respiratory distress) and muscles (resulting in infarcts or muscle pain). Fever, tachycardia, tachypnoea, anorexia and depression are also frequently present. Renal dysfunction is common.

In horses with evidence of PH, haematological and plasma biochemical analysis may reveal a leukopaenia or leukocytosis, anaemia, thrombocytopaenia, hyperfibrinogenaemia, hypo‐ or normoproteinaemia and increased concentrations of muscle enzymes. A presumptive diagnosis is generally made based on the history, clinical signs and exclusion of other causes of oedema. Elevated titres of IgA and IgG against S. equi are supportive of a diagnosis. Very high SeM‐specific antibody titres (>1:12,800) are strongly suggestive of PH secondary to strangles infection or vaccination. Skin biopsies can confirm leukocytoclastic vasculitis but are rarely performed.

The objective of treatment is to reduce inflammation. Tapering doses of corticosteroids are used – typically dexamethasone, with an initial dose of 0.1–0.2 mg/kg IM or IV for 3–5 days and then reducing the dose by half every 3–5 days over 2–4 weeks. Antimicrobials may be necessary where bacterial infection is concurrent or considered likely. Other supportive care such as hydrotherapy, supportive bandaging, IV fluids and non‐steroidal anti‐inflammatory drugs are usually required.