WHAT'S THE POINT?
Intravenous (IV) cannulation for the administration of fluids and medications is probably the most commonly performed invasive procedure in small animal veterinary practice but can be uncomfortable for the patient and stressful for both the animal and clinic staff (Chebroux et al, 2015). In this article we will examine the reasons for IV cannulation, cannula design, commonly used locations, placement techniques and how to minimise frequently encountered complications.
Most commonly, peripheral intravenous (IV) cannulation is used for the administration of fluids and medications in the peri-anaesthetic period or for the management of medical cases. Short-term intravenous cannulation optimises the speed and efficacy of drug, fluid, blood product and parenteral nutrition administration (Beal & Hughes, 2000), reduces the incidence of thrombus formation, thrombophlebitis, occlusion, sepsis and air emboli (Brismar & Nystrom, 1986; Karapinar & Cura, 2007; Nickel, 2019), and provides a means for blood sample collection (Tan et al, 2003). Highly irritant fluids e.g. chemotherapy agents, may be administered via IV cannulae to reduce the possibility of extravascular deposition, tissue irritation and/or necrosis (Crow, 1997). Peripheral IV cannulation offers improved patient welfare by avoiding the requirement for repeated injections. The treatment of anaesthetic emergencies is greatly facilitated by the presence of a patent IV cannula.
The most commonly used material for veterinary peripheral intravenous cannulae (PIVC) is polyurethane although they can also be made from Teflon or silicon. Polyurethane is flexible and biostable and minimises stress on vessel walls, which in turn reduces the potential for thrombophlebitis and sepsis (Spurlock et al, 1990).
The majority if PIVC used in veterinary medicine are “over the needle” and come in a variety of gauges, lengths and designs. The inner metal stylet and cannula are introduced into the vein, the cannula advanced and the stylet removed. For small animal practice 18G-24G are the most widely used although 14, 16 and 26G are also available.
Guide-wire catheters are also obtainable and are useful for hypovolaemic patients with difficult to locate veins, and for neonates.
The ideal PIVC should occupy no more than 50-60% of the vein diameter e.g. a 22G cannula with an external diameter of 0.9mm is suitable for vein diameter of approximately 1.8mm.
A PIVC that is too wide may result in obstruction to blood flow and restricted movement of administered medications/fluids. The overlarge PIVC can also abrade vessel walls increasing the possibility of phlebitis and rupture (the “blown” vein).
If the cannula is too small for the vein inadequate flow of medications e.g. fluids, may occur, and pressure alarms on infusion devices can be triggered. Withdrawing blood from inadequately sized PIVCs can also cause iatrogenic damage to erythrocytes (Braun, 2018).
The species and breed of patient can influence the site selected for IVC placement. Some breeds e.g. brachycephalics, Dachshunds, may have limb conformities that make venous visualisation and access difficult (Goddard, 2010).
The most commonly accessed vessels for PIVC are the cephalic and saphenous veins although the marginal ear vein (particularly in rabbits) and jugular are also commonly used (Tan et al, 2003).
The cephalic vein is easy to visualise and access but care should be taken to ensure the tip of the cannula is located distal to the elbow joint as flexion may reduce blood flow and patency of the cannula.
In the hind leg the lateral saphenous vein is used most commonly. Although readily visualised the vein is mobile and this may present challenges during catheter insertion and maintenance (Tan et al, 2003). In patients experiencing diarrhoea, excessive urination or urinary/faecal incontinence there is a high risk of hind limb catheter site complications and the cephalic vein is preferable for PIVC placement (Goddard, 2010).
The size of the jugular vein allows placement of a large gauge cannula for high volume fluid administration. The vessel is easily located although in some short-necked breeds the function of the cannula may be impaired, and many patients do not tolerate the manipulation and dressings required in this location (Tan et al, 2003)
Patient preparation, hand hygiene and aseptic technique are all essential during PIVC placement to minimise the risk of infection (Ashby, 2017; Goddard, 2010). Skin and hair can be a source of infection, so preparation should be as diligent as for a surgical procedure (Crow, 1997).
- All equipment should be prepared and easily available. Avoid placing equipment and materiasl on the floor (if the animal has to be restrained in such a location) or any contaminated surface (Goddard, 2010).
- The patient should be gently restrained in a clean, pre-prepared location.
- Whilst preparing the catheter site, non-sterile gloves should ideally be worn following thorough hand washing or use of an alcohol hand rub (McMillan & Ackerman, 2016).
- A cannulation site distant from a joint and free from skin damage or infection should be selected.
- To reduce sepsis and improve visualisation & cannula security clippers should be used to remove sufficient hair to prevent contamination of the connection system (Goddard, 2010). Shaving is associated with increased infection rates and should be avoided (Tanner et al, 2011).
- An assistant raises the vein and it is assessed for suitability.
- The site is prepared as for routine surgery. Young et al (2014) suggest the use of 2% chlorhexidine in 70% isopropyl alcohol for site preparation although Scarlet (2012) advocates the use of 2% chlorhexidine followed by isopropyl alcohol. Chlorhexidine has been associated with lower infection rates than povidone iodine (Scarlet, 2012). In a study by Dorey-Phillips & Murrison (2008) chlorhexidine was found to be more effective than industrial methylated spirits for reducing bacterial counts on canine skin.
- Hand preparation should be repeated and new gloves donned.
- The vein is raised and the tip of the cannula, bevel side up, is inserted into the vein at an ideal angle of 10-30o (Braun, 2018). A flash of blood will be seen in the hub of the cannula.
- The stylet is held stationary whilst the cannula is advanced into the vein. Do not advance the stylet.
- Care should be taken to avoid contamination of the outer surface of the cannula during the entire insertion process.
- The stylet is removed.
- The assistant releases the vein. Pressure may be applied over the vein to reduce haemorrhage.
- The selected connector/bung is attached.
- The cannula is flushed with sterile 0.9% saline or heparinised saline to ensure patency (may be performed after securing the cannula with tape).
- The PIVC is secured in place with tape and dressings. These should be selected based on the anticipated duration of cannulation, the procedure and predicted degree handling.
- Tape tension should be monitored. Limb swelling may occur if the dressing is too tight. (Taylor et al, 2011).
- Further dressings may be applied to minimise gross contamination of the PIVC site.
- The presence of a protective dressing can prevent observation of the PIVC site during administration of medications therefore these should ideally be removed during treatment to allow assessment of cannula patency/connector security and to monitor for extravasation of fluids/drugs/other medications.
PIVC placement can be uncomfortable and distressing for the patient therefore gentle, thoughtful handling is necessary and the use of EMLA cream or other form of local anaesthesia should be considered. In a 2018 study by van Oostrom & Knowles the use of EMLA cream applied 60 minutes prior to intravenous cannulation in dogs reduced patient discomfort.
Maintenance of the IV cannula
Dressings and taping should be monitored regularly – if too tight distal limb oedema may occur, and if too loose the dressings can dislodge the PIVC (Weil, 2006). If the cannula is not being used for the administration of fluids or constant rate infusions the insertion site should be assessed at least daily, ideally more frequently, for signs of infection, inflammation or catheter displacement (Goddard, 2013; Taylor et al, 2011) and the PIVC removed if any abnormal observations are made.
Dressings should be changed if they become wet or soiled and this will provide an opportunity to also examine the cannulation site.
PIVC patency should be checked by regular flushing (Taylor et al, 2011). There is some variation between authors on the recommended flush frequency: Weil (2006) advises every 2-4 hours whereas Scarlet (2012) suggests every 2-6 hours. Either heparinised saline or 0.9% sodium chloride may be used as the flushing fluid: Davis (2015) concluded they were both equally effective. The patient should be monitored during PIVC flushing procedures for signs of discomfort or pain as this may indicate dislodgement. If discomfort is observed the cannula should be examined carefully and removed if there is any suspicion of misplacement (Taylor et al, 2011).
Drug administration and flushing
Following the administration of any drug, fluid or blood product via a PIVC the catheter and associated bung/port/tap/line should be flushed with sterile (heparinised) normal saline. Many drugs are irritant and if present at high concentrations within the cannula can cause vessel inflammation. Additionally, the cannula and bung/port/tap can accommodate a significant volume of fluid and failure to flush can result in underdosing/overdosing the patient, particularly for low volume/high potency drugs and/or in small patients. Macfie (1990) calculated that 10-30% of a 1ml injection could remain in the cannula. The consequences of not flushing following drug administration were demonstrated by Singleton et al (2005): late onset muscle paralysis occurred in patients where succinyl choline remaining in equipment dead space was injected following subsequent fluid administration. This study also reported precipitation in equipment dead space when 2 incompatible drugs were sequentially administered without adequate flushing between products.
- Extravascular positioning and subsequent perivascular administration of drugs is relatively common. Removal of protective dressings, so that the cannula insertion site and surrounding tissues are visible during drug administration, will allow early detection of a “blown” vein.
- Inadvertent mixing of incompatible drugs can occur if the cannula and bung are not flushed thoroughly (Singleton et al, 2005).
- Under- or over-dosing may occur if the cannula and bung/connector are not flushed (Singleton, 2005).
- Infection is relatively common. In animals not demonstrating any signs of cannula infection 19% of the IV cannulae had positive bacterial cultures, 19.7% of cannulae used for blood sample collection immediately after insertion tested positive, and 8.3% of cannulae attached to Y or T connectors had positive cultures (Jones et al, 2009). In the same study animals with signs of local infection produced 42.9%, 34.8% and 21.1% positive cultures from the cannula itself, following blood sampling via the cannula, and from catheters attached to Y/T connectors respectively. Skin organisms were most frequently cultured suggesting particular attention to strict aseptic technique is important to reduce the incidence of infection.
Intravenous cannulation is a common invasive procedure in veterinary practice and should be performed in all patients undergoing sedation or anaesthesia. Attention should be paid to strict asepsis during insertion to reduce the possibility of infection and chlorhexidine is more effective than alcohol alone for reducing cannula site infections. EMLA cream many improve patient comfort during insertion. Cannulae and insertion sites should be regularly inspected for infection, patency and positioning. Removal of bulky dressings will aid detection of extravasation and the cannula & bung/connector should be thoroughly flushed following drug administration.
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Originally published: Thursday, 2nd May 2019
Ashby J. (2017). Peripheral intravenous catheter care in hospitalised cats and dogs. Vet Nursing J. 32: 32-36
Beal M.W. & Hughes D. (2000). Vascular access: theory and techniques in the small animal emergency patient. Clin Tech Small An Prac. 15: 101-109
B. Braun, Australia. (2018). Choosing the correct IV catheter. www.bbraun.com.au.
Chebroux S, Leece E.A. & Brearley J.C. (2015). Ease of intravenous catheterisation in dogs and cats: a comparative study of two peripheral catheters. JSAP. 56: 242-246
Crow S.E. (1997). Placement and care of intravenous catheters. In: Manual of clinical procedures in the dog, cat and rabbit. 2nd ed. Lippincott. 45-66
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Goddard L. (2010). How to obtain vascular access: the importance of good placement and aseptic technique. Vet Nurse. 1: 50-53
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Scarlet F. (2012). Small animal anaesthesia and the role of the nurse: Part 3. Vet Times, June 2012.
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