• An arterial blood sample is used to measure the arterial O2 tension (PaO2), CO2 tension (PaCO2), pH and bicarbonate/base excess levels, and Hb saturation (SaO2).
• Familiarize yourself with the location and use of the blood gas machine. Arterial blood is obtained either by percutaneous needle puncture or from an indwelling arterial line.
• Radial artery: more accessible and more comfortable for the patient; best palpated between the bony head of the distal radius and the tendon of the flexor carpi radialis, with the wrist dorsiflexed. The Allen test is used to identify impaired collateral circulation in the hand (a contraindication to radial artery puncture)—the patient’s hand is held high, with the fist clenched, and both the radial and ulnar arteries are compressed. The hand is lowered, the fist is opened, and the pressure from the ulnar artery is released. Colour should return to the hand within 5s.
• Brachial artery: best palpated medial to the biceps tendon in the antecubital fossa, with the arm extended and the palm facing up. The needle is inserted just above the elbow crease.
• Femoral artery: best palpated just below the midpoint of the inguinal ligament, with the leg extended. The needle is inserted below the inguinal ligament at a 90° angle.
• The chosen puncture site should be cleaned. Local anaesthetic should be infiltrated (not into the artery). Use one hand to palpate the artery, and the other hand to advance the heparin-coated syringe and needle (22–25G) at a 60–90° angle to the skin, with gentle aspiration. A flush of bright red blood indicates successful puncture. Remove about 2–3mL of blood; withdraw the needle, and ask an assistant to apply pressure to the puncture site for 5–15min. Air bubbles should be removed. The sample is placed on ice and analysed within 15min (to reduce O2 consumption by WBC).
• Complications: include persistent bleeding, bruising, injury to the blood vessel, and local thrombosis.
• Continuous monitoring of arterial BP in critically ill patients with haemodynamic instability.
• Repeated arterial blood sampling.
• Raynaud’s phenomenon.
• Thromboangiitis obliterans.
• Advanced atherosclerosis.
• End-arteries, such as the brachial artery, should be avoided.
• Locate a palpable artery (e.g. radial or femoral).
• Assess the ulnar blood flow using the Allen test before inserting a radial line ( Arterial blood sampling, p. [link]).
• Position the hand in moderate dorsiflexion, with the palm facing up (to bring the artery closer to the skin).
• The site should be cleaned with a sterile preparation solution and draped appropriately.
• Use sterile gloves.
• Use local anaesthetic (1% lidocaine) in a conscious patient.
(See Fig. 15.1.)
• Palpate the artery with the non-dominant hand (1–2cm from the wrist, between the bony head of the distal radius and the flexor carpi radialis tendon).
• The catheter and needle are then advanced through the vessel a few millimetres further (see Fig. 15.1c).
• The needle is removed (see Fig. 15.1d).
• The catheter is slowly withdrawn until pulsatile blood flow is seen (see Fig. 15.1e).
• When pulsatile blood flow is seen, the wire is advanced into the vessel (see Fig. 15.1f).
• The catheter is advanced further into the vessel over the wire (see Fig. 15.1g).
• While placing pressure over the artery, the wire is removed (see Fig. 15.1h) and the catheter is connected to a transduction system.
• Secure the catheter in place using suture or tape.
• Check perfusion to the hand after insertion of the arterial line and at frequent intervals.
• The line should be removed if there are any signs of vascular compromise or as early as possible after it is no longer needed.
(See Fig. 15.2.)
Locate and palpate the artery with the non-dominant hand (1–2cm from the wrist, between the bony head of the distal radius and the flexor carpi radialis tendon).
• The catheter and needle are then advanced slightly further, and the catheter/needle angle is lowered to 10–15° (see Fig. 15.2c).
• The catheter is advanced over the needle into the vessel (see Fig. 15.2d).
• Proximal pressure is applied to the artery; the needle is removed (see Fig. 15.2e) and the catheter is connected to a transduction system.
• Secure the catheter in place using suture or tape.
• Check perfusion to the hand after insertion of the arterial line and at frequent intervals.
• Local and systemic infection.
• Bleeding, haematoma, bruising.
• Vascular complications: blood vessel injury, pseudoaneurysm, thromboembolism, and vasospasm.
• Arterial spasm may occur after multiple unsuccessful attempts at arterial catheterization. If this occurs, use an alternative site.
• There may be difficulty in passing a wire or catheter, despite the return of pulsatile blood. Adjustment of the angle, withdrawal of the needle, or a slight advance may be helpful.
You will need
• Sterile dressing pack, gloves, and sterile occlusive dressing.
• A 5- and 10-mL syringe, green (21G) and orange (25G) needles.
• Local anaesthetic (e.g. 2% lidocaine), saline flush.
• Central line (e.g. 16G long Abbocath® or Seldinger catheter).
• Silk suture and needle, No. 11 scalpel blade.
• Arterial puncture (remove and apply local pressure).
• Pneumothorax (insert a chest drain or aspirate if required).
• Haemothorax or chylothorax (mainly left subclavian lines).
• Infection (local, septicaemia, bacterial endocarditis).
• Brachial plexus or cervical root damage (infiltration with local anaesthetic).
• The basic technique is the same whatever vein is cannulated.
• Lie the patient supine (± head-down tilt).
• Turn the patient’s head away from the side you wish to use.
• Clean the skin with chlorhexidine: from the angle of the jaw to the clavicle for IJV cannulation, and from the midline to the axilla for the subclavian approach.
• Use drapes to isolate the sterile field.
• Flush the lumen of the central line with saline.
• Infiltrate the skin and subcutaneous tissue with local anaesthetic.
• Have the introducer needle and the Seldinger guidewire within easy reach, so that you can reach them with one hand without having to release your other hand. Your fingers may be distorting the anatomy slightly, making access to the vein easier and, if released, it may prove difficult to relocate the vein.
• With the introducer needle in the vein, check that you can aspirate blood freely. Use the hand that was on the pulse to immobilize the needle relative to the skin.
• Remove the syringe and pass the guidewire into the vein; it should pass freely. If there is resistance, remove the wire; check that the needle is still within the lumen, and try again.
• Remove the needle, leaving the wire within the vein, and use a sterile swab to maintain gentle pressure over the site of venepuncture to prevent excessive bleeding.
• With a No. 11 blade, make a nick in the skin where the wire enters, to facilitate dilatation of the subcutaneous tissues. Pass the dilator over the wire and remove, leaving wire in situ.
• Pass the central line over the wire into the vein. Remove the guidewire; flush the lumen with fresh saline, and close to air.
• Suture the line in place, and cover the skin penetration site with a sterile occlusive dressing.
• Measuring the CVP (see Box 15.1).
The IJV runs just posterolateral to the carotid artery within the carotid sheath and lies medial to the sternocleidomastoid (SCM) in the upper part of the neck, between the two heads of the SCM in its medial portion, and enters the subclavian vein (SCV) near the medial border of the anterior scalene muscle (see Fig. 15.3a). There are three basic approaches to IJV cannulation: medial to the SCM, between the two heads of the SCM, or lateral to the SCM. The approach used varies and depends on the experience of the operator and the institution.
• Locate the carotid artery between the sternal and clavicular heads of the SCM at the level of the thyroid cartilage; the IJV lies just lateral and parallel to it.
• Keeping the fingers of one hand on the carotid pulsation, infiltrate the skin with local anaesthetic thoroughly, aiming just lateral to this and ensuring that you are not in a vein.
• Ideally, first locate the vein with a blue or green needle. Advance the needle at 45° to the skin, with gentle negative suction on the syringe, aiming for the ipsilateral nipple, lateral to the pulse.
• If you fail to find the vein, withdraw the needle slowly, maintaining negative suction on the syringe (you may have inadvertently transfixed the vein). Aim slightly more medially and try again.
• Once you have identified the position of the vein, change to the syringe with the introducer needle; cannulate the vein, and pass the guidewire into the vein (see Fig. 15.3).
Tips and pitfalls
• Venous blood is dark, and arterial blood is pulsatile and bright red!
• Once you locate the vein, change to the syringe with the introducer needle, taking care not to release your fingers from the pulse; they may be distorting the anatomy slightly, making access to the vein easier and, if released, it may prove difficult to relocate the vein.
• The guidewire should pass freely down the needle and into the vein. With the left IJV approach, there are several acute bends that need to be negotiated. If the guidewire keeps passing down the wrong route, ask your assistant to hold the patient’s arms out at 90° to the bed, or even above the patient’s head, to coax the guidewire down the correct path.
• For patients who are intubated or requiring respiratory support, it may be difficult to access the head of the bed. The anterior approach may be easier (see Fig. 15.3b) and may be done from the side of the bed (the left side of the bed for right-handed operators, using the left hand to locate the pulse and the right to cannulate the vein).
• The IJV may also be readily cannulated with a long Abbocath®. No guidewire is necessary, but, as a result, misplacement is more common than with the Seldinger technique.
• When using an Abbocath®, on cannulating the vein, remember to advance the sheath and needle a few millimetres to allow the tip of the plastic sheath (~1mm behind the tip of the bevelled needle) to enter the vein. Holding the needle stationary, advance the sheath over it into the vein.
• Arrange for a CXR to confirm the position of the line.
The axillary vein becomes the SCV at the lateral border of the first rib and extends for 3–4cm just deep to the clavicle. It is joined by the ipsilateral IJV to become the brachiocephalic vein behind the sternoclavicular joint. The subclavian artery and brachial plexus lie posteriorly, separated from the vein by the scalenus anterior muscle. The phrenic nerve and the internal mammary artery lie behind the medial portion of the SCV and, on the left, lies the thoracic duct (see Fig. 15.4).
• Select the point 1cm below the junction of the medial third and middle third of the clavicle. If possible, place a bag of saline between the scapulae to extend the spine.
• Clean the skin with iodine or chlorhexidine.
• Infiltrate the skin and subcutaneous tissue and the periosteum of the inferior border of the clavicle with local anaesthetic up to the hilt of the green (21G) needle, ensuring that it is not in a vein.
• Insert the introducer needle with a 10-mL syringe, guiding gently under the clavicle. It is safest to initially hit the clavicle, and ‘walk’ the needle under it until the inferior border is just cleared. In this way, you keep the needle as superficial to the dome of the pleura as possible. Once it has skimmed underneath the clavicle, advance it slowly towards the contralateral sternoclavicular joint, aspirating as you advance. This technique minimizes the risk of pneumothorax, with high success.
• Once venous blood is obtained, rotate the bevel of the needle towards the heart. This encourages the guidewire to pass down the brachiocephalic vein, rather than up the IJV.
• The wire should pass easily into the vein. If there is difficulty, try advancing during the inspiratory and expiratory phases of the respiratory cycle.
• Once the guidewire is in place, remove the introducer needle, and make a small incision in the skin near the wire to allow the dilator to pass over the wire. When removing the dilator, note the direction that it faces; it should be slightly curved downwards. If it is slightly curved upwards, then it is likely that the wire has passed up into the IJV. When this happens, it is safer to remove the wire and start again.
• After removing the dilator, pass the central venous catheter over the guidewire; remove the guidewire, and secure.
• A CXR is mandatory after subclavian line insertion, to exclude a pneumothorax and to confirm satisfactory placement of the line, especially if fluoroscopy was not employed.
Traditional central venous catheterization methods rely on anatomical landmarks to predict vein position. However, the relationship between such landmarks and the vein position varies significantly in ‘normal’ individuals. Failure and complication rates using landmark methods are significant, and therefore serious complications may occur. Recent advances in portable US equipment have now made it possible to insert central venous catheters under 2D US guidance.
Advantages of this technique include:
• Identification of actual and relative vein positions.
• Identification of anatomical variations.
• Confirmation of target vein patency.
Guidelines from NICE (September 2002) state: ‘Two-dimensional imaging ultrasound guidance is recommended as the preferred method for insertion of central venous catheters into the internal-jugular vein (IJV) in adults and children in elective situations’. However, training and equipment availability render such recommendations effectively useless in the UK at present.
• Standard Seldinger-type kit or whatever is locally available.
• An assistant is essential.
• US equipment:
• Screen: displays 2D US image of anatomical structures.
• Sheaths: dedicated sterile sheaths of polyvinyl chloride (PVC) or latex long enough to cover the probe and connecting cable (a rubber band secures the sheath to the probe).
• Probe: a transducer which emits and receives US information to be processed for display. Marked with an arrow or a notch for orientation.
• Power: battery or mains.
• Sterile gel: transmits US and provides a good interface between the patient and the probe.
Sterile precautions should be taken, with the patient’s head turned slightly away from the cannulation site. Head-down tilt (if tolerated) or leg elevation to increase filling and the size of the IJV. Ensure adequate drapes to maintain a sterile field.
Excessive head rotation or extension may decrease the diameter of the vein.
• Ensure that the display can be seen.
• The sheath is opened (operator) and the gel squirted in (assistant). A generous amount of gel ensures good contact and air-free coupling between the probe tip and the sheath. Too little may compromise the image quality.
• The probe and connecting cable are lowered into the sheath (assistant), which is then unrolled along them (operator).
• A rubber band secures the sheath to the probe.
• The sheath over the probe tip is smoothed out (wrinkles will degrade the image quality).
• Apply liberal amounts of gel to the sheathed probe tip for good US transmission and patient comfort during movement.
The most popular scanning orientation for IJV central catheter placement is the transverse plane.
• Apply the probe tip gently to the neck, lateral to the carotid pulse, at the cricoid level, or in the sternomastoid–clavicular triangle.
• Keep the probe perpendicular at all times, with the tip flat against the skin.
• Orientate the probe so that movement to the left ensures that the display looks to the left (and vice versa). Probes are usually marked to help orientation. By convention, the mark should be to the patient’s right (transverse plane) or to the head (longitudinal scan). The marked side appears on the screen as a bright dot.
• If the vessels are not immediately visible, keep the probe perpendicular and gently glide medially or laterally until found.
When moving the probe, watch the screen—not your hands.
After identification of the IJV
• Position the probe so that the IJV is shown at the display’s horizontal midpoint.
• Keep the probe immobile.
• Direct the needle (bevel towards probe) caudally under the marked midpoint of the probe tip at ~60° to the skin.
• The needle bevel faces the probe to help direct the guidewire down the IJV later.
• Advance the needle towards the IJV.
Needle passage causes a ‘wavefront’ of tissue compression. This is used to judge the progress of the needle and position. Absence of visible tissue reaction indicates incorrect needle placement. Just before vessel entry, ‘tenting’ of the vein is usually observed.
One of the most difficult aspects to learn initially is the steep needle angulation required, but this ensures that the needle enters the IJV in the US beam and takes the shortest and most direct route through the tissues.
Needle pressure may oppose vein walls, resulting in vein transfixion. Slow withdrawal of the needle with continuous aspiration can help result in lumen access.
Pass the guidewire into the jugular vein in the usual fashion.
Re-angling the needle from 60° to a shallower angle, e.g. 45°, may help guidewire feeding. Scanning the vein in the longitudinal plane may demonstrate the catheter in the vessel, but after securing and dressing the central venous catheter, an X-ray should still be obtained to confirm the central venous catheter position and exclude pneumothorax.
The most common error in measurement of the CVP, particularly in CVP lines which have been in place for some time, is due to partial or complete line blockade. With the manometer connected, ensure that the line is free-flowing; minor blockages can be removed by squeezing the rubber bung, with the line proximal being obliterated by acute angulation (i.e. bend the tube proximal). Measure the CVP at the mid-axillary line, with the patient supine. CVP falls with upright or semi-upright recumbency, regardless of the reference point. If the CVP is high, lift the stand that holds the manometer so that the apparent CVP falls by 10cm or so, and re-place the CVP stand to ground level. If the saline or manometer reading rises to the same level, then the CVP reading is accurate. In other words, one ensures that the CVP manometer level both falls and rises to the same level.
PA catheters (Swan–Ganz catheters) allow direct measurement of a number of haemodynamic parameters that aid clinical decision-making in critically ill patients (evaluate RV and LV function, guide treatment, and provide prognostic information). The catheter itself has no therapeutic benefit, and there have been a number of studies showing mortality (and morbidity) with their use. Consider inserting a PA catheter in any critically ill patient, after discussion with an experienced physician, if the measurements will influence decisions on therapy (and not just to reassure yourself). Careful and frequent clinical assessment of the patient should always accompany measurements, and PA catheterization should not delay treatment of the patient.
General indications (not a comprehensive list) include:
• Management of complicated MI.
• Assessment and management of shock.
• Assessment and management of respiratory distress (cardiogenic versus non-cardiogenic pulmonary oedema).
• Evaluating effects of treatment in unstable patients (e.g. inotropes, vasodilators, mechanical ventilation, etc.).
• Delivering therapy (e.g. thrombolysis for PE, epoprostenol for pulmonary hypertension, etc.).
• Assessment of fluid requirements in critically ill patients.
• Full resuscitation facilities should be available, and the patient’s ECG should be continuously monitored.
• Bag of heparinized saline for flushing the catheter and a transducer set for pressure monitoring. (Check that your assistant is experienced in setting up the transducer system before you start.)
• An 8F introducer kit (prepackaged kits contain the introducer sheath and all the equipment required for central venous cannulation).
• PA catheter: commonly a triple-lumen catheter, that allows simultaneous measurement of RA pressure (proximal port) and PA pressure (distal port) and incorporates a thermistor for measurement of cardiac output by thermodilution. Check your catheter before you start.
• Fluoroscopy is preferable, though not essential.
• Do not attempt this, unless you are experienced.
• Observe a strict aseptic technique using sterile drapes, etc.
• Insert the introducer sheath (at least 8F in size) into either the IJV or the SCV in the standard way. Flush the sheath with saline, and secure to the skin with sutures.
• Do not attach the plastic sterile expandable sheath to the introducer yet, but keep it sterile for use later once the catheter is in position (the catheter is easier to manipulate without the plastic covering).
• Flush all the lumens of the PA catheter, and attach the distal lumen to the pressure transducer. Check the transducer is zeroed (conventionally to the mid-axillary point). Check the integrity of the balloon by inflating it with the syringe provided (2mL of air), and then deflate the balloon.
(See Fig. 15.5.)
Pulmonary artery catheterization 2
• Flush all the lumens of the PA catheter, and attach the distal lumen to the pressure transducer. Check the transducer is zeroed (conventionally to the mid-axillary point). Check the integrity of the balloon by inflating it with the syringe provided (~2mL of air), and then deflate the balloon.
• Pass the tip of the PA catheter through the plastic sheath, keeping the sheath compressed. The catheter is easier to manipulate without the sheath over it; once in position, extend the sheath over the catheter to keep it sterile.
• With the balloon deflated, advance the tip of the catheter to ~10–15cm from the right IJV or SCV, 15–20cm from the left (the markings on the side of the catheter are at 10cm intervals: two lines = 20cm). Check that the pressure tracing is typical of the RA pressure (see Fig. 15.6 and Table 15.1).
• Inflate the balloon and advance the catheter gently. The flow of blood will carry the balloon (and catheter) across the tricuspid valve, through the RV, and into the PA.
• Watch the ECG tracing closely, while the catheter is advanced. The catheter commonly triggers runs of VT when crossing the tricuspid valve and through the RV. VT is usually self-limiting but should not be ignored. Deflate the balloon, pull back, and try again.
• If >15cm of the catheter is advanced into the RV without the tip entering the PA, this suggests the catheter is coiling in the RV. Deflate the balloon, withdraw the catheter into the RA, reinflate the balloon, and try again using clockwise torque while advancing in the ventricle or flushing the catheter with cold saline to stiffen the plastic. If this fails repeatedly, try under fluoroscopic guidance.
• As the tip passes into a distal branch of the PA, the balloon will impact and not pass further; the wedge position and pressure tracing will change (see Fig. 15.6).
• Deflate the balloon, and check that a typical PA tracing is obtained. If not, try flushing the catheter lumen, and, if that fails, withdraw the catheter until the tip is within the PA, and begin again.
• Reinflate the balloon slowly. If the PCWP is seen before the balloon is fully inflated, it suggests the tip has migrated further into the artery. Deflate the balloon and withdraw the catheter 1–2cm, and try again.
• If the pressure tracing flattens and then continues to rise, you have ‘overwedged’. Deflate the balloon, pull back the catheter 1–2cm, and start again.
• When a stable position has been achieved, extend the plastic sheath over the catheter and secure it to the introducer sheath. Clean any blood from the skin insertion site with antiseptic, and secure a coil of the PA catheter to the patient’s chest to avoid inadvertent removal.
• Obtain a CXR to check the position of the catheter. The tip of the catheter should ideally be no more than 3–5cm from the midline.
Table 15.1 Normal values of right heart pressures and flows
Right atrial pressure
Pulmonary capillary wedge pressure
Tips and pitfalls
• Never withdraw the catheter with the balloon inflated.
• Never advance the catheter with the balloon deflated.
• Never inject liquid into the balloon.
• Never leave the catheter with the balloon inflated, as pulmonary infarction may occur.
• The plastic of the catheter softens with time at body temperature, and the tip of the catheter may migrate further into the PA branch. If the pressure tracing with the balloon deflated is ‘partially wedged’ (and flushing the catheter does not improve this), withdraw the catheter 1–2cm and reposition.
• Sometimes it is impossible to obtain a wedged trace. In this situation, one has to use the PA diastolic pressure as a guide. In health, there is 72–4mmHg difference between the PA diastolic pressure and PCWP. Any condition which causes pulmonary hypertension (e.g. severe lung disease, ARDS, long-standing valvular disease) will alter this relationship.
• Valvular lesions, VSDs, prosthetic valves, and pacemakers: if these are present, then seek advice from a cardiologist. The risk of subacute bacterial endocarditis (SBE) may be sufficiently great that the placement of a PA catheter may be more detrimental than beneficial.
• PEEP ( Positive end-expiratory pressure, p. [link]) measurement and interpretation if PCWP in patients on PEEP depends on the position of the catheter. Ensure the catheter is below the level of the LA on a lateral CXR. Removing PEEP during measurement causes marked fluctuations in haemodynamics and oxygenation, and the pressures do not reflect the state once back on the ventilator.
• Arrhythmias: watch the ECG tracing closely, while the catheter is advanced. The catheter commonly triggers runs of VT when crossing the tricuspid valve and through the RV. If this happens, deflate the balloon, pull back, and try again. VT is usually self-limiting but should not be ignored.
• PA rupture (~0.2% in one series): damage may occur if the balloon is overinflated in a small branch. Risk factors include MV disease (large v wave confused with poor wedging), pulmonary hypertension, multiple inflations, or hyperinflations of the balloon. Haemoptysis is an early sign. It is safer to follow PA diastolic pressures if these correlate with the PCWP.
• Pulmonary infarction.
• Knots: usually occur at the time of initial placement in patients where there has been difficulty in traversing the RV. Signs include loss of pressure tracing, persistent ectopy, and resistance to catheter manipulation. If this is suspected or has occurred, stop manipulation and seek expert help.
• Infection: risks increase with the length of time the catheter is left in situ. The pressure transducer may occasionally be a source of infection. Remove the catheter and introducer, and replace only if necessary.
• Other complications: complications associated with central line insertion, thrombosis and embolism, balloon rupture, and intracardiac damage.
1 Following acute MI
• Symptomatic CHB (any territory).
• Symptomatic secondary heart block (any territory).
• Trifascicular block:
• Alternating LBBB and RBBB.
• First-degree heart block + RBBB + left axis deviation.
• New RBBB and left posterior hemiblock.
• LBBB and long PR interval.
• After anterior MI:
• Asymptomatic CHB.
• Asymptomatic second-degree (Mobitz II) block.
• Symptomatic sinus bradycardia unresponsive to atropine.
• Recurrent VT for atrial or ventricular overdrive pacing.
2 Unrelated to MI
• Symptomatic sinus or junctional bradycardia unresponsive to atropine (e.g. carotid sinus hypersensitivity).
• Symptomatic secondary heart block or sinus arrest.
• Symptomatic CHB.
• Torsades de pointes tachycardia.
• Recurrent VT for atrial or ventricular overdrive pacing.
• Bradycardia-dependent tachycardia.
• Drug OD (e.g. verapamil, β-blockers, digoxin).
• Permanent pacemaker box change in a patient who is pacing-dependent.
3 Before general anaesthesia
• The same principles as for acute MI (see earlier).
• Sinoatrial disease and secondary (Wenckebach) heart block only need prophylactic pacing if there are symptoms of syncope or pre-syncope.
Transvenous temporary pacing
• The most commonly used pacing mode and the mode of choice for life-threatening bradyarrhythmias is ventricular demand pacing (VVI) with a single bipolar wire positioned in the RV (see Temporary cardiac pacing: ventricular pacing, pp. [link]–[link] for an explanation of common pacing modes).
• In critically ill patients with impaired cardiac pump function and symptomatic bradycardia (especially with RV infarction), cardiac output may be by up to 20% by maintaining AV synchrony. This requires two pacing leads, one atrial and one ventricular, and a dual pacing box.
Following cardiac surgery, patients may have epicardial wires (attached to the pericardial surface of the heart) left in for up to 1 week in case of post-operative heart block or bradyarrhythmia. These may be used in the same way as the more familiar transvenous pacing wires, but the threshold may be higher.
AV sequential pacing
In critically ill patients with impaired cardiac pump function and symptomatic bradycardia (especially with RV infarction), cardiac output may be by up to 20% by maintaining AV synchrony. This requires two pacing leads, one atrial and one ventricular, and a dual pacing box.
• Cannulate a central vein: the wire is easiest to manipulate via the right internal jugular (RIJ) approach but is more comfortable for the patient via the right SCV. The left internal jugular (LIJ) approach is best avoided, as there are many acute bends to negotiate and a stable position is difficult to achieve. Avoid the left subclavicular area, as this is the preferred area for permanent pacemaker insertion and should be kept ‘virgin’, if possible. The femoral vein may be used, but the risk of DVT and infection is high.
• Insert a sheath: (similar to that for PA catheterization) through which the pacing wire can be fed. Pacing wires are commonly 5F or 6F, and a sheath at least one size larger is necessary. Most commercially available pacing wires are prepacked with an introducer needle and a plastic cannula similar to an Abbocath® which may be used to position the pacing wire. However, the cannula does not have a haemostatic seal. The plastic cannula may be removed from the vein, leaving the bare wire entering the skin, once a stable position has been achieved. This reduces the risk of wire displacement but also makes repositioning of the wire more difficult, should this be necessary, and the infection risk is higher.
• Pass the wire through the sterile plastic cover that accompanies the introducer sheath, and advance into the upper RA (see Fig. 15.7), but do not unfurl the cover yet. The wire is much easier to manipulate with gloved hands, without the additional hindrance of the plastic cover.
• Advance the wire, with the tip pointing towards the RV; it may cross the tricuspid valve easily. If it fails to cross, point the tip to the lateral wall of the atrium and form a loop. Rotate the wire, and the loop should fall across the tricuspid valve into the ventricle.
• Advance and rotate the wire, so that the tip points inferiorly as close to the tip of the RV (laterally) as possible.
• If the wire does not rotate down to the apex easily, it may be because you are in the coronary sinus, rather than in the RV. (The tip of the wire points to the left shoulder.) Withdraw the wire, and re-cross the tricuspid valve.
• Leave some slack in the wire; the final appearance should be like the outline of a sock, with the ‘heel’ in the RA, the ‘arch’ over the tricuspid valve, and the ‘big toe’ at the tip of the RV.
• Connect the wire to the pacing box, and check the threshold. Ventricular pacing thresholds should be <1.0V, but a threshold of up to 1.5V is acceptable if another stable position cannot be achieved.
• Check for positional stability. With the box pacing at a rate higher than the intrinsic HR, ask the patient to take some deep breaths, cough forcefully, and sniff. Watch for failure of capture, and, if so, reposition the wire.
• Set the output to 3V and the box on ‘demand’. If the patient is in sinus rhythm and has an adequate BP, set the box rate to just below the patient’s rate. If there is CHB or bradycardia, set the rate at 70–80/min.
• Cover the wire with the plastic sheath, and suture the sheath and wire securely to the skin. Loop the rest of the wire, and fix to the patient’s skin with adhesive dressing.
• When the patient returns to the ward, obtain a CXR to confirm satisfactory positioning of the wire and to exclude a pneumothorax.
(See Fig. 15.8.)
• Advance the atrial wire until the ‘J’ is reformed in the RA.
• Rotate the wire, and withdraw slightly to position the tip in the RA appendage. Aim for a threshold of <1.5V.
• If atrial wires are not available, a ventricular pacing wire may be manipulated into a similar position or passed into the coronary sinus for LA pacing.
(See Box 15.3.)
Ventricular ectopics or VT
• Non-sustained VT is common, as the wire crosses the tricuspid valve (especially in patients receiving an isoprenaline infusion), and does not require treatment.
• Try to avoid long runs of VT and, if necessary, withdraw the wire into the atrium and wait until the rhythm has settled.
• If ectopics persist after the wire is positioned, try adjusting the amount of slack in the wire in the region of the tricuspid valve (either more or less).
• Pacing the RVOT can provoke runs of VT.
Failure to pace and/or sense
• It is difficult to get low pacing thresholds (<1.0V) in patients with extensive MI (especially of the inferior wall) or cardiomyopathy or who have received class I antiarrhythmic drugs. Accept a slightly higher value if the position is otherwise stable and satisfactory.
• If the position of the wire appears satisfactory and yet the pacing thresholds are high, the wire may be in a left hepatic vein. Pull the wire back into the atrium and try again, looking specifically for ventricular ectopics as the wire crosses the tricuspid valve.
• The pacing threshold commonly doubles in the first few days due to endocardial oedema.
• If the pacemaker suddenly fails, the most common reason is usually wire displacement:
• Increase the pacing output of the box.
• Check all the connections of the wire and the battery of the box.
• Try moving the patient to the left lateral position until arrangements can be made to reposition the wire.
• A pericardial rub may be present in the absence of perforation (especially post-MI).
• Presentation: pericardial chest pain, increasing breathlessness, falling BP, enlarged cardiac silhouette on CXR, signs of cardiac tamponade, left diaphragmatic pacing at low output.
• High-output pacing (10V), even with a satisfactory position of the ventricular lead, may cause pacing of the left hemidiaphragm. At low voltages, this suggests perforation (see Perforation, p. [link]).
• Right hemidiaphragm pacing may be seen with atrial pacing and stimulation of the right phrenic nerve.
• Reposition the wire if symptomatic (painful twitching, dyspnoea).
Establish peripheral venous access, and check that full facilities for resuscitation are available. Pre-prepared pericardiocentesis sets may be available. You will need:
• A trolley, as for central line insertion, with iodine or chlorhexidine for the skin, dressing pack, sterile drapes, local anaesthetic (lidocaine 2%), syringes (including a 50mL), needles (25G and 22G), a No. 11 blade, and silk sutures.
• Pericardiocentesis needle (15cm, 18G) or similar Wallace cannula.
• J-guidewire (≥80cm, 0.035in diameter).
• Dilators (up to 7F).
• Pigtail catheter (≥60cm with multiple sideholes, a large Seldinger-type CVP line can be used if no pigtail is available).
• Drainage bag and connectors.
• Facilities for fluoroscopy or echocardiographic screening.
(See Fig. 15.9.)
• Position the patient at ~30°. This allows the effusion to pool inferiorly within the pericardium.
• Sedate the patient lightly with midazolam and fentanyl if necessary. Use with caution, as this may drop the BP in patients already compromised by the effusion.
• Put on a sterile gown and gloves; clean the skin from mid chest to mid abdomen, and place the sterile drapes on the patient.
• Infiltrate the skin and subcutaneous tissues with local anaesthetic, starting 1–1.5cm below the xiphisternum and just to the left of the midline, aiming for the left shoulder and staying as close to the inferior border of the rib cartilages as possible.
• The pericardiocentesis needle is introduced into the angle between the xiphisternum and the left costal margin, angled at >30°. Advance slowly, aspirating gently and then injecting more lidocaine every few millimetres, aiming for the left shoulder.
• As the parietal pericardium is pierced, you may feel a ‘give’ and fluid will be aspirated. Remove the syringe, and introduce the guidewire through the needle.
• Check the position of the guidewire by screening. It should loop within the cardiac silhouette only and not advance into the SVC or PA.
• Remove the needle, leaving the wire in place. Enlarge the skin incision slightly, using the blade, and dilate the track.
• Insert the pigtail over the wire into the pericardial space, and remove the wire.
• Take specimens for microscopy, culture (and inoculate a sample into blood culture bottles), cytology, and haematocrit if bloodstained (an FBC tube; ask the haematologists to run on a Coulter counter for a rapid estimation of Hb).
• Aspirate to dryness, watching the patient carefully. Symptoms and haemodynamics (tachycardia) often start to improve with removal of as little as 100mL of pericardial fluid.
• If the fluid is heavily bloodstained, withdraw fluid cautiously; if the pigtail is in the RV, withdrawal of blood may cause cardiovascular collapse. Arrange for urgent Hb/haematocrit.
• Leave on free drainage and attached to the drainage bag.
• Suture the pigtail to the skin securely, and cover with a sterile occlusive dressing.
• Closely observe the patient for recurrent tamponade (obstruction of the drain), and repeat Echo.
• Discontinue anticoagulants.
• Remove the drain after 24h or when drainage stops.
• Consider the need for surgery (drainage, biopsy, or pericardial window) or specific therapy (chemotherapy if malignant effusion, antimicrobials if bacterial, dialysis if renal failure, etc.).
See Box 15.4 for complications of pericardiocentesis.
Tips and pitfalls
If the needle touches the heart’s epicardial surface
You may feel a ‘ticking’ sensation transmitted down the needle—withdraw the needle a few millimetres; angulate the needle more superficially, and try cautiously again, aspirating as you advance.
If you do not enter the effusion
• Withdraw the needle slightly and advance again, aiming slightly deeper, but still towards the left shoulder.
• If this fails, try again, aiming more medially (mid-clavicular point or even suprasternal notch).
• Consider trying the apical approach (starting laterally at the cardiac apex and aiming for the right shoulder) if Echo confirms sufficient fluid at the cardiac apex.
Difficulty in inserting the pigtail
• This may be because of insufficient dilatation of the tract.
• Hold the wire tort (by gentle traction), while pushing the catheter; take care not to pull the wire out of the pericardium.
Haemorrhagic effusion versus blood
• Compare the Hb of the pericardial fluid to venous blood Hb.
• Place some of the fluid in a clean container; blood will clot, whereas a haemorrhagic effusion will not, as the ‘whipping’ action of the heart tends to defibrinate it.
• Confirm the position of the needle by first withdrawing some fluid and then injecting 10–20mL of contrast; using fluoroscopy, see if the contrast stays within the cardiac silhouette.
• Alternatively, if using Echo guidance, inject 5–10mL of saline into the needle, looking for ‘microbubble contrast’ in the cavity containing the needle tip. Injecting 20mL of saline rapidly into a peripheral vein will produce ‘contrast’ in the RA and RV and may allow them to be distinguished from the pericardial space.
• Connect a pressure line to the needle; a characteristic waveform will confirm penetration of the RV (see Fig. 15.6).
• Digoxin toxicity.
• Electrolyte disturbance (Na+, K+, Ca2+, Mg2+, acidosis).
• Inadequate anticoagulation and chronic AF.
See Box 15.5 for complications of DC cardioversion.
Checklist for DC cardioversion
Check this is functioning, with a fully equipped arrest trolley to hand in case of an arrest.
(Unless life-threatening emergency.)
AF, flutter, SVT, VT, signs of ischaemia or digoxin. If the ventricular rate is slow, have an external (transcutaneous) pacing system nearby in case of asystole.
For at least 4h.
Does the patient require anticoagulants? Is the INR >2.0? (Has it been so for >3 weeks?)
Check this is >3.5mmol/L.
Check there are no features of digoxin toxicity and recent digoxin levels are normal. If there are frequent ventricular ectopics, give IV Mg2+ 8mmol.
Treat thyrotoxicosis or myxoedema first.
Peripheral venous cannula.
Short general anaesthesia (propofol) is preferable to sedation with benzodiazepine and fentanyl. Bag the patient with 100% O2.
See Table 15.2.
Check this is selected on the defibrillator for all shocks (unless the patient is in VF or haemodynamically unstable). Adjust the ECG gain, so that the machine is only sensing QRS complexes, and not P or T waves.
Most centres now use ‘hands-free’ adhesive paddles for DC cardioversion. Some continue with the traditional handheld paddles.
Conductive gel pads should be placed between the right of the sternum and the other to the left of the left nipple (anterior to mid-axillary line). Alternatively, place one anteriorly just left of the sternum, and one posteriorly to the left of the midline. There is some evidence that the anteroposterior (AP) position is superior for AF.
Check no one is in contact with the patient or with the metal bed. Ensure your own legs are clear of the bed! Apply firm pressure on the paddles if using the handheld device.
Double the energy level, and repeat up to 360J.
Consider changing the paddle position (see ‘Paddle placement’ earlier in the table). If prolonged sinus pause or ventricular arrhythmia during an elective procedure, stop.
• When complete, repeat ECG. Place in the recovery position until awake. Monitor for 2–4h, and ensure the effects of sedation have passed. Patients should be accompanied home by a friend or relative if being discharged.
Table 15.2 Suggested initial energies for DC shock for elective cardioversion
• If the initial shock is unsuccessful, increase the energy (50, 100, 200, 360J) and repeat.
• If still unsuccessful, consider changing the paddle position and try 360J again. It is inappropriate to persist further with elective DC cardioversion.
The risk of thromboembolism in patients with chronic AF and dilated cardiomyopathy is 0–7%, depending on the underlying risk factors.
• Age <60 years.
• No heart disease.
• Recent-onset AF (<3 days).
Anticoagulate patients at risk with warfarin for at least 3–4 weeks. For recent-onset AF (1–3 days), anticoagulate with IV heparin for at least 12–24h and, if possible, exclude an intracardiac thrombus with TOE prior to DC shock. If there is a thrombus, anticoagulate with warfarin, as described earlier. For emergency cardioversion of AF (<24h), heparinize prior to shock.
The risk of systemic embolism with cardioversion of atrial flutter and other tachyarrhythmias is very low, provided there is no ventricular thrombus, since the coordinated atrial activity prevents the formation of a clot. Routine anticoagulation with warfarin is not necessary, but we would recommend heparin before DC shock, as the atria are often rendered mechanically stationary for several hours after shock, even though there is coordinate electrical depolarization.
After successful cardioversion, if the patient is on warfarin, continue anticoagulation for at least 3–4 weeks. Consider indefinite anticoagulation if there is intrinsic cardiac disease (e.g. mitral stenosis) or recurrent AF.
DC shock during pregnancy appears to be safe. Auscultate the fetal heart before and after cardioversion and, if possible, the fetal ECG should be monitored.
There is a danger of damage to the pacemaker generator box or the junction at the tip of the pacing wire(s) and endocardium. Position the paddles in the AP position, as this is theoretically safer. Facilities for backup pacing (external or transvenous) should be available. Check the pacemaker post-cardioversion—both early and late problems have been reported.
• Cardiogenic shock post-MI.
• Acute severe MR.
• Acute VSD.
• Preoperative (ostial left coronary stenosis).
• Weaning from cardiopulmonary bypass.
• Dilated cardiomyopathy.
• Aortic dissection.
• Severe aorto-iliac atheroma.
• Bleeding diathesis.
• Aortic dissection.
• Arterial perforation
• Peripheral embolism.
• Limb ischaemia.
• Balloon rupture.
The device consists of a catheter with a balloon (40mL size) at its tip which is positioned in the descending thoracic aorta. Balloon inflation/deflation is synchronized to the ECG. The balloon should inflate just after the dicrotic notch (in diastole), thereby increasing pressure in the aortic root and increasing coronary perfusion. The balloon deflates just before ventricular systole, thereby decreasing afterload and improving LV performance (see Fig. 15.10).
Counterpulsation has many beneficial effects on the circulation:
• coronary perfusion in diastole.
• Reduced LVEDP.
• Reduced myocardial O2 consumption.
• cerebral and peripheral blood flow.
Previous experience is essential. Formerly, a cut-down to the femoral artery was required, but newer balloons come equipped with a sheath which may be introduced percutaneously. Using fluoroscopy, the balloon is positioned in the descending thoracic aorta, with the tip just below the origin of the left subclavian artery. Fully anticoagulate the patient with IV heparin. Some units routinely give IV antibiotics (flucloxacillin) to cover against Staphylococcus infection.
Triggering and timing
The balloon pump may be triggered either from the patient’s ECG (R wave) or from the arterial pressure waveform. Slide switches on the pump allow precise timing of inflation and deflation during the cardiac cycle. Set the pump to 1:2 to allow you to see the effects of augmentation on alternate beats.
• Seek help from an expert! There is usually an on-call cardiac perfusionist or technician, a senior cardiac physician, or a surgeon.
• Counterpulsation is inefficient with HRs over 130bpm. Consider antiarrhythmics or 1:2 augmentation instead.
• Triggering and timing: for ECG triggering, select a lead with the most pronounced R wave; ensure that the pump is set to trigger from the ECG, not pressure; permanent pacemakers may interfere with triggering-select lead with negative and smallest pacing artefacts. Alternatively, set the pump to be triggered from the external pacing device. A good arterial waveform is required for pressure triggering; the timing will vary slightly, depending on the location of the arterial line (slightly earlier for radial artery line, cf. femoral artery line). Be guided by the haemodynamic effects of balloon inflation and deflation, rather than the precise value of delay.
• Limb ischaemia: exacerbated by poor cardiac output, adrenaline, NA, and peripheral vascular disease. Wean off and remove the balloon (see IABP removal, p. [link]).
• Thrombocytopenia: commonly seen; does not require transfusion, unless there is overt bleeding, and returns to normal once the balloon is removed. Consider epoprostenol infusion if platelet counts fall below 100 × 109/L.
• The patient may be progressively weaned by gradually reducing the counterpulsation ratio (1:2, 1:4, 1:8, etc.) and/or reducing the balloon volume and checking that the patient remains haemodynamically stable.
• Stop the heparin infusion, and wait for the ACT to fall <150s (APTT <1.5 normal).
• Using a 50mL syringe, have an assistant apply negative pressure to the balloon.
• Pull the balloon down until it abuts the sheath; do not attempt to pull the balloon into the sheath.
• Withdraw both balloon and sheath, and apply firm pressure on the femoral puncture site for at least 30min or until bleeding is controlled.
The aim of therapy is to relieve hypoxia and maintain or restore a normal PaCO2 for the individual. Relative indications for mechanical ventilation are discussed in the appropriate chapters. This section discusses some of the principles involved.
• O2 should be administered by a system that delivers a defined percentage, between 28% and 100%, according to the patient’s requirements (e.g. via fixed-percentage delivery masks such as Ventimask Mk IV).
• A Hudson mask or nasal cannulae give very variable FiO2, depending on the flow rate and the patient’s breathing pattern.
• Nasal prongs only deliver at FiO2 of 30% at flows of 2L/min, and become less efficient at higher flow rates (>35% at 3L/min, with little further increase with increasing flow). Higher flow rates require humidification.
• A properly positioned high-flow O2 mask, using O2 at 6L/min, can provide an FiO2 of 60%.
• Combining nasal prongs and a high-flow mask can achieve an FiO2 of >80–90%.
• In practice, it is rarely possible to consistently deliver >60%, unless using CPAP or ventilation.
• When sudden deterioration in oxygenation occurs, check the delivery system for empty cylinders, disconnected tubing, etc.
• Type 1 or 2 respiratory failure.
• Bronchial asthma.
• Acute MI.
• Sickle-cell crisis.
• CO poisoning.
• Cluster headaches.
• Tracheobronchitis occurs with prolonged inhalation of ≥80% O2. It causes retrosternal pain, cough, and dyspnoea.
• Parenchymal lung damage from O2 occurs with FiO2 >60% for >48h without intermittent air breathing periods.
Monitoring oxygen therapy
• O2 therapy should be assessed by continuous oximetry and intermittent ABGs.
• Oximetry is an invaluable aid but has limitations. In some situations (e.g. GBS), falling oximetry is a very late marker of impending respiratory failure, and CO2 accumulation (e.g. in COPD) is clearly not monitored by oximetry. An SaO2 of 93% correlates with a PaO2 of 8kPa, and below 92%, the PaO2 may fall disproportionately quickly.
• Periodic ‘sighs’ are a normal part of breathing and reverse microatelectasis. Lung expansion techniques are indicated for patients who cannot or will not take periodic large breaths (e.g. post-abdominal or chest surgery, neuromuscular chest wall weakness).
• Post-operative techniques used commonly by physiotherapists include incentive spirometry, coached maximal inspiration with cough, postural drainage, and chest percussion.
• Volume-generating devices, such as ‘the Bird’, are triggered by the patient initiating inspiration and deliver a preset tidal volume to augment the patient’s breath. Liaise with your physiotherapist.
• Pressure-generating techniques [such as CPAP, nasal intermittent positive pressure ventilation (NIPPV), and bilevel positive airway pressure (BiPAP)] have the advantage that, even if a leak develops around the mask, the ventilator is able to ‘compensate’ to provide the prescribed positive pressure (see following sections).
• For both volume- and pressure-generating techniques, the patient must be able to protect their airway and generate enough effort to trigger the machine.
Continuous positive airways pressure
• CPAP provides a constant positive pressure throughout the respiratory cycle.
• It acts to splint open collapsing alveoli which may be full of fluid (or a collapsing upper airway in OSA), increases functional residual capacity (FRC) and compliance, such that the work of breathing is reduced and gas exchange is improved.
• It allows a higher FiO2 (approaching 80–100%) to be administered, cf. standard O2 delivery masks.
• CPAP should usually be commenced after liaison with anaesthetists; in a patient for active management, it should usually be started on the ITU.
• A standard starting pressure is 5cmH2O.
• Pulmonary oedema.
• Acute respiratory failure (e.g. secondary to infection) where a simple face mask for O2 is insufficient.
• Acute respiratory failure where ventilation is inappropriate.
• Weaning from the ventilator.
• Patient needs to:
• Be awake and alert.
• Be able to protect the airway.
• Possess adequate respiratory muscle strength.
• Be haemodynamically stable.
Negative pressure ventilation (NPV)
• This works by ‘sucking’ out the chest wall and is used in chronic hypoventilation (e.g. polio, kyphoscoliosis, or muscle disease). Expiration is passive.
• These techniques do not require tracheal intubation. However, access to the patient for nursing care is difficult.
Intermittent positive pressure ventilation (IPPV)
Deteriorating gas exchange due to a potentially reversible cause of respiratory failure:
• Head injury.
• Exacerbation of COPD.
• Cerebral hypoxia.
• Massive atelectasis (e.g. post-cardiac arrest).
• Respiratory muscle weakness.
• Intracranial bleed.
• Myasthenia gravis.
• Raised ICP.
• Acute infective polyneuritis.
• Major trauma or burns.
Ventilation of the ill patient on the ITU is via either an ETT or a tracheostomy. If ventilation is anticipated to be needed for >1 week, consider a tracheostomy.
There are two basic types of ventilator.
• Pressure-cycled ventilators deliver gas into the lungs until a prescribed pressure is reached, when inspiratory flow stops and, after a short pause, expiration occurs by passive recoil. This has the advantage of reducing the peak airway pressures without impairing cardiac performance in situations such as ARDS. However, if the airway pressures increase or compliance decreases, the tidal volume will fall, so patients need to be monitored closely to avoid hypoventilation.
• Volume-cycled ventilators deliver a preset tidal volume into the lungs over a predetermined inspiratory time (usually ~30% of the breathing cycle), hold the breath in the lungs (for ~10% of the cycle), and then allow passive expiration as the lungs recoil.
• NIPPV delivers positive pressure for a prescribed inspiratory time, when triggered by the patient initiating a breath, allowing the patient to exhale to atmospheric pressure.
• The positive pressure is supplied by a small machine via a tight-fitting nasal mask.
• It is generally used as a method of home nocturnal ventilation for patients with severe musculoskeletal chest wall disease (e.g. kyphoscoliosis) or with OSA.
• It has also been used, with modest success, as an alternative to formal ventilation via ETT in patients where positive expiratory pressure is not desirable, e.g. acute asthma, COPD with CO2 retention, and as a weaning aid in those in whom separation from a ventilator is proving difficult.
• The system is relatively easy to set up by experienced personnel, but some patients take to it better than others. It should not be commenced by inexperienced personnel.
Continuous mandatory ventilation
• Continuous mandatory ventilation (CMV) acts on a preset cycle to deliver a given number of breaths per minute of a set volume. The duration of the cycle determines the breath frequency.
• The minute volume is calculated by (tidal volume × frequency).
• The relative proportions of time spent in inspiration and expiration (I:E ratio) is normally set at 1:2 but may be altered. For example, in acute asthma, where air trapping is a problem, a longer expiratory time is needed ( Acute severe asthma: further management, p. [link]); in ARDS, where the lung compliance is low, a longer inspiratory time is beneficial (inverse ratio ventilation; Adult respiratory distress syndrome 2, p. [link]).
• The patients should be fully sedated. Patients capable of spontaneous breaths who are ventilated on CMV can get ‘stacking’ of breaths where the ventilator working on its preset cycle may give a breath on top of one which the patient has just taken, leading to overinflation of the lungs, a high peak inspiratory pressure, and the risk of pneumothorax.
• Prolonged use of this mode will result in atrophy of the respiratory muscles; this may prove difficult in subsequent ‘weaning’, especially in combination with proximal myopathy from steroids, e.g. in acute asthma.
• Ventilation may either be terminated abruptly or by gradual transfer of the ventilatory workload from the machine to the patient (‘weaning’).
Synchronized intermittent mandatory ventilation
• Synchronized intermittent mandatory ventilation (SIMV) modes allow the patient to breath spontaneously and be effectively ventilated, and allows gradual transfer of the work of breathing on to the patient. This may be appropriate when weaning the patient whose respiratory muscles have wasted. It is inappropriate in acutely ill patients (e.g. acute severe asthma, ARDS); CMV with sedation reduces O2 requirement and respiratory drive, and allows more effective ventilation.
• Exact details of the methods of synchronization vary between machines, but all act in a similar manner—the patient breathes spontaneously through the ventilator circuit. The ventilator is usually preset to ensure that the patient has a minimum number of breaths per minute, and if the number of spontaneous breaths falls below the preset level, then a breath is delivered by the machine.
• Most SIMV modes of ventilation provide some form of positive pressure support to the patient’s spontaneous breaths, to reduce the work of breathing and ensure effective ventilation (see Pressure support, p. [link]).
• Positive pressure is added during inspiration to relieve part or all of the work of breathing.
• This may be done in conjunction with an SIMV mode of ventilation or as a means of supporting entirely spontaneous patient-triggered ventilation during the process of weaning.
• It allows the patients to determine their own RR and should ensure adequate inflation of the lungs and oxygenation. It is, however, only suitable for those whose lung function is reasonably adequate and who are not confused or exhausted.
Positive end-expiratory pressure
• PEEP is a preset pressure added to the end of expiration only, to maintain the lung volume, prevent airway or alveolar collapse, and open up atelectatic or fluid-filled lungs (e.g. in ARDS or cardiogenic pulmonary oedema).
• It can significantly improve oxygenation by making more of the lung available for gas exchange. However, the trade-off is an increase in intrathoracic pressure, which can significantly decrease venous return and hence cardiac output. There is also an risk of pneumothorax.
• ‘Auto-PEEP’ is seen if the patient’s lungs do not fully empty before the next inflation (e.g. asthma).
• In general, PEEP should be kept at a level of 5–10cmH2O, where required, and the level adjusted in 2–3cmH2O intervals every 20–30min, according to a balance between oxygenation and cardiac performance.
• Measurement and interpretation of PCWP in patients on PEEP depend on the position of the catheter. PCWP will always reflect pulmonary venous pressures if they are greater than PEEP. If the catheter is in an apical vessel where the PCWP is normally lower, due to the effects of gravity, the pressure measured may be the alveolar (PEEP) pressure, rather than the true PCWP; in a dependent area, the pressures are more accurate. Removing PEEP during measurement alters the haemodynamics and oxygenation, and the pressures do not reflect the state once back on the ventilator.
• To bypass upper airway obstruction (e.g. trauma, infections, neoplasms, post-operative, burns, and corrosives) when oral or nasotracheal intubation is contraindicated.
• In situations when ET intubations fail (e.g. massive nasopharyngeal haemorrhage, structural deformities, obstruction due to foreign bodies, etc.).
The Seldinger technique is quicker, may be performed by non-surgeons at the bedside, and is safer (see Fig. 15.11). After anaesthetizing the area, a needle is used to puncture the cricothyroid membrane, and, through this, a guidewire is introduced into the trachea. Over this, a series of dilators and a tracheostomy tube can be safely positioned.
This is the best method for providing and maintaining a clear airway for ventilation, protection against aspiration, and suctioning and clearing lower respiratory tract secretions. The most common indication for urgent intubation by a physician is cardiac arrest. This is not a technique for the inexperienced—the description given here is not intended as a substitute for practice under supervision of a skilled anaesthetist.
You will need
• Laryngoscope, usually with a curved blade (Macintosh).
• ETT (8–9mm internal diameter for ♂ and 7–8mm for ♀) and appropriate adaptors.
• Syringe for cuff inflation, and clamp to prevent air escaping from the cuff once inflated.
• Scissors and tape or bandage to secure the tube.
• Lubricating jelly (e.g. K-Y® jelly).
• Suction apparatus with rigid (Yankauer) and long, flexible catheters.
Potential problems during intubation
• Certain anatomical variations (e.g. receding mandible, short neck, prominent incisors, high-arched palate), as well as stiff neck or trismus, may make intubation complicated; summon experienced help.
• Vomiting: suction if necessary. Cricoid pressure may be useful.
• Cervical spine injury: immobilize the head and neck in line with the body, and try not to extend the head during intubation.
• Facial burns or trauma may make orotracheal intubation impossible. Consider cricothyrotomy ( Percutaneous cricothyrotomy, p. [link]).
(See Fig. 15.12.)
• Place the patient with the neck slightly flexed and the head extended. Take care if cervical injury is suspected.
• Cricoid pressure: the oesophagus can be occluded by compressing the cricoid cartilage posteriorly against the body of C6. This prevents passive regurgitation into the trachea, but not active vomiting. Ask your assistant to maintain pressure until the tube is in place and the cuff inflated.
• Pre-oxygenate the patient by hyperventilation with ≥85% O2 for 15–30s. Suction the throat to clear the airway.
• With the laryngoscope in your left hand, insert the blade on the right side of the mouth. Advance to the base of the tongue, identifying the tonsillar fossa and the uvula. Push the blade to the left, moving the tongue over. Advance the blade until the epiglottis comes into view.
• Insert the blade tip between the base of the tongue and the epiglottis, and pull the whole blade (and larynx) upwards along the line of the handle of the laryngoscope to expose the vocal cords. Brief suction may be necessary to clear the view.
• If the cords cannot be seen, do not poke at the epiglottis, hoping for success; call for more skilled help, and revert to basic airway management.
• Intubation must not take longer than 30s; if there is any doubt about the position, remove the tube, reoxygenate, and try again.
• With the tube in place, listen to the chest during inflation to check that both sides of the chest inflate. If the tube is in the oesophagus, chest expansion will be minimal, though the stomach may inflate.
• Tie the ETT in place to prevent it from slipping up or down the airway. Ventilate with high-concentration O2.
If the pneumothorax is <75% and the patient is haemodynamically stable, it is reasonable to attempt aspiration of the pneumothorax in the first instance ( Pneumothorax: assessment, pp. [link]–[link]).
You will need
• 10mL and 50mL syringes with green (18G) and orange (25G) needles.
• Dressing pack (swabs, sterile drapes, antiseptic) and sterile gloves.
• 19G Venflon® or alternative cannula.
• Local anaesthetic (e.g. 2% lidocaine).
• A 3-way tap.
• One assistant is required.
• Sit the patient up, propped against pillows, with their hand behind their head; ensure you are comfortable and on a similar level.
• Select the space to aspirate—the second intercostal space in the mid- clavicular line. Confirm with a CXR that you are aspirating the correct side (a surprisingly common cause of disasters is aspirating the normal side).
• Clean the skin and use an aseptic technique.
• Connect a 50mL syringe to a 3-way tap in readiness, with the line which will be connected to the patient turned ‘off’ so that no air will enter the pleural cavity on connecting the apparatus.
• Infiltrate 5–10mL of lidocaine from the skin to pleura, just above the upper border of the rib in the space you are using. Confirm the presence of air by aspirating ~5mL via a green needle.
• Insert a 16G or larger IV cannula into the pneumothorax, preferably while aspirating the cannula with a syringe, so that entry into the pleural space is confirmed. Allow the tip of the cannula to enter the space by ~1cm.
• Ask the patient to hold their breath, and remove the needle. Swiftly connect the 3-way tap. Aspirate 50mL of air/fluid, and void it through the other lumen of the tap. Repeat.
• Aspiration should be stopped when resistance to suction is felt, the patient coughs excessively, or ≥2.5L of air has been aspirated.
• Withdraw the cannula, and cover the site with a dressing plaster (e.g. Elastoplast™ or Band-Aid™)
• Check a post-procedure CXR. If there is significant residual pneumothorax, insert a chest drain.
The basic procedure is similar to that for a pneumothorax; the site is different—one or two intercostal spaces posteriorly below the level at which dullness is detected. Ideally, all cases should have an USS first to confirm the level of the effusion and ensure that the diaphragm is not higher than anticipated due to underlying pulmonary collapse.
• Position the patient leaning forward over the back of a chair or table. Clean the skin, and infiltrate with local anaesthetic, as described for a pneumothorax aspiration.
• Insert the cannula, and aspirate the effusion with a 50mL syringe, voiding it through the 3-way tap. Repeat until resistance is felt and the tap is dry.
• Check a post-procedure CXR.
You will need
• Dressing pack (sterile gauze, gloves, drapes, povidone-iodine).
• Local anaesthetic (720mL of 1% lidocaine), 10mL syringe, green (18G) and orange (25G) needles.
• Scalpel and No. 11 blade for skin incision; two packs of silk sutures (1–0).
• Two forceps (Kelly clamps), scissors, needle-holder (often prepackaged as a ‘chest drain set’).
• Where possible, use the new Seldinger-type chest tubes—especially for pneumothorax.
• Chest tubes—a selection of 24, 28, 32, and 36F.
• Chest drainage bottles, with sterile water for underwater seal.
• One assistant.
• Position the patient leaning forward over the back of a chair or table. If possible, premedicate the patient with an appropriate amount of opiate ~30min before.
• Mark the space to be drained in the mid-axillary line—usually the fifth intercostal space for a pneumothorax, and below the level of the fluid for an effusion. Clean the skin.
• Select the chest tube: small (24F) for air alone, medium (28F) for serous fluid, or large (32–36F) for blood/pus. Remove the trocar. Check that the underwater seal bottles are ready.
• Infiltrate the skin with 15–20mL of lidocaine 1%. Make a short subcutaneous tunnel for the chest tube before it enters the pleural space (see Fig. 15.13). Anaesthetize the periosteum on the top of the rib. Check that you can aspirate air/fluid from the pleural space.
• Make a horizontal 2cm incision in the anaesthetized skin of the rib space. Use the forceps to blunt-dissect through the fat and intercostal muscles to make a track large enough for your gloved finger down to the pleural space. Stay close to the upper border of the rib to avoid the neurovascular bundle.
• Check the length of the tube against the patient’s chest to confirm how much needs to be inserted into the patient’s chest. Aim to get the tip to the apex for a pneumothorax; keep the lowermost hole as low as possible (>2cm into the chest) to drain pleural fluid.
• Insert two sutures across the incision (or a purse-string, see Fig. 15.13). These will gently tighten around the tube, once inserted, to create an airtight seal, but do not knot—these sutures will be used to close the wound after drain removal.
• Remove the trocar. Clamp the end of the tube with the forceps, and gently introduce the tube into the pleural space. Rotating the forceps 180° directs the tube to the apex (see Fig. 15.13). Condensation in the tube (or fluid) confirms the tube is within the pleural space. Check that all the holes are within the thorax and connect to the underwater seal. Tape these to the skin.
• Gently tighten the skin sutures, but do not knot. The drain should be secured with several other stitches and copious amounts of adhesive tape. They are very vulnerable to accidental traction.
• Wrap adhesive tape around the join between the drain and the connecting tubing.
• Prescribe adequate analgesia for the patient for when the local anaesthetic wears off.
• Arrange for a CXR to check the position of the drain.
• Do not drain off >1L of pleural fluid/24h to avoid re-expansion pulmonary oedema.
Tips and pitfalls
• The chest drain should only be left in place, while air or fluid continues to drain. The risk of an ascending infection increases with time. Prophylactic antibiotics are not usually indicated.
• Obtain a CXR (and then daily) to check the position of the drain and examine the lung fields.
• If the drain is too far out, there will be an air leak and the patient may develop subcutaneous emphysema. Ideally, remove the drain and replace with a new drain at a new site; the risk of an ascending infection is high if the ‘non-sterile’ portion of the tube is just pushed into the chest.
• If the drain is too far in, it may be uncomfortable for the patient and impinge on vital structures (e.g. thoracic aorta). Pull the tube out the appropriate distance and re-suture.
• Check the water column in the chest drain bottle swings with respiration. This stops if the tube is obstructed.
• Check the drains and tubing are free of bends and kinks.
• Blood clots or fibrin may block the tube.
• If the lung is still collapsed on the CXR, replace the chest drain with a new tube at a new site.
Lung fails to re-expand
• This is either due to an obstructed system or a persistent air leak (e.g. tracheobronchial fistula).
• If the chest drain continues to bubble, apply suction to the drain to help expand the lung. Consider inserting further drains or surgical repair of the leak. If the chest drain is obstructed (described earlier), replace the drain.
Removing the chest drain
• Do not clamp the chest drain.
• Remove the dressings, and release the sutures holding the drain in place. Leave the skin incision sutures (purse-string) in position to close the wound once the drain is removed.
• Remove the drain in a gentle motion, either in inspiration or in expiration with Valsalva.
• Tighten the skin sutures. These should be removed after 3–4 days and a fresh dressing applied.
• Any residual pneumothorax should be treated, depending on the patient’s symptoms.
• To diagnose or exclude SBP.
• To obtain ascites for measurement of protein, albumin, or amylase (pancreatic ascites).
• Ascitic cytology may require 100mL of fluid.
• Stain and culture for AFBs; lymphocytes: >500 cells/mm3).
• To drain cirrhotic or malignant ascites.
• Previous abdominal surgery increases the risk of perforation (due to adhesion of underlying bowel to the abdominal wall).
• Massive hepatomegaly or splenomegaly (avoid the same side).
• Massive ileus with bowel distension.
NB There are no clinical data to support avoiding paracentesis in severe coagulopathy (platelets <20 000, INR >4.0), but most clinicians should be cautious and consider correcting coagulopathy.
• Lie the patient supine and tilted slightly to one side.
• Select the site for paracentesis (e.g. on a horizontal line across the umbilicus, and 4cm lateral to a line passing to the mid-inguinal point). Clean the area with chlorhexidine. Avoid surgical scars (see Fig. 15.14).
• Use a 20mL syringe with a 18G (green) needle. In obese patients, use a longer needle (e.g. 18G Abbocath®). Infiltrate the area with local anaesthetic. Insert the needle slowly into the abdomen, while aspirating until fluid is obtained.
• Inoculate 5mL of the fluid into each bottle of a set of blood culture bottles, and send 5mL in a sterile bottle for microscopy and protein determination. Add 2mL of ascites to an EDTA tube (contains anticoagulant), and send to haematology for cell count.
• Remove and apply a sterile plaster over the puncture site.
Daily small-volume paracentesis increases the risk of complications such as infection and ascitic leakage. The risk of infection is high if a peritoneal drain is left in situ in cirrhotic ascites. It is safer to drain the ascites to dryness.
The rate of fluid drainage can be fast, and it is generally safe to drain >3–5L/h. During the first 3–6h of paracentesis, there is a significant increase in cardiac output, a decrease in SVR, and a modest fall in MAP (by 5–10mmHg). Tense ascites increases the RA pressure, which falls acutely following paracentesis.
• Previous abdominal surgery and scarring. Be cautious—avoid scars, and use USS guidance.
• Patients with clinically apparent DIC or fibrinolysis and oozing from needle-sticks.
• Paracentesis should not be performed in patients with a massive ileus with bowel distension without imaging guidance.
• Platelet count <20. Ignore the INR (FFP not required).
• Renal failure: the risk of bleeding is in renal failure, although this is NOT a contraindication, and diagnosis of infected ascites is important in this group of patients.
• It is important to avoid puncturing the superficial inferior epigastric artery, which runs just lateral to the umbilicus from the mid-inguinal point (see Fig. 15.14). Avoid any visible superficial veins.
• For USS-guided paracentesis, mark the spot prior to the procedure.
• The skin should be cleaned and sterilized. Use chlorhexidine pads. Anaesthetize the area (see ‘X’ in Fig. 15.14) with a small volume of lidocaine.
• Do NOT go too lateral, and the patient should be supine and leaning slightly to one side.
• Beware of patients with a very large spleen or very large liver, and avoid going too close to either structure. Use USS guidance, if possible.
• To avoid the catheter blocking due to omentum plugging the end, use a catheter with side-holes.
Insertion of drainage catheter and draining the ascites
• You will need to gown up with a sterile gown.
• Monitor the BP before, and hourly for the first 6h.
• Insert the drainage catheter with multiple side-holes provided (attached to a 20mL syringe), aspirating as one advances the cannula. When ascitic fluid is obtained, advance the needle 3–4cm more, and then advance the plastic cannula into the abdomen and attach the drainage system.
• Ask the patient to lie to one side, so that the drainage side is downmost (left or right).
• Allow all ascites to drain as quickly as it will come, regardless of the volume. When the ascites stops draining or slows down, move the patient from side to side and lie towards the drainage site.
• When complete, remove the catheter; apply plaster, and lie the patient with the drainage site uppermost for at least 4h (~100mL of 20% albumin for every 2.5L removed).
• Replace albumin with an infusion of 20% albumin to give 8g of albumin for every litre of ascites removed. It is usually best to start the albumin infusion after paracentesis is complete, but if there is a significant drop in BP, it can be started earlier.
• Try and ensure that all fluid is drained, as incomplete paracentesis increases the risk of leakage post-procedure.
• Measure the volume of ascites drained.
• Always remove the drainage catheter within 6h of insertion to decrease the risk of sepsis.
The Sengstaken–Blakemore tube is inserted to control variceal bleeding when endoscopic therapy or IV terlipressin have failed. It should not be used as primary therapy, since it is unpleasant and increases the risk of oesophageal ulceration and aspiration.
Seek experienced or specialist help early. Balloon tamponade is a temporary procedure to prevent exsanguination.
• It is assumed that the patient is undergoing resuscitation and has received IV terlipressin. To reduce the risk of aspiration, the patient should be intubated and ventilated.
• The Sengstaken tube should be stored in the fridge (to maximize stiffness) and removed just before use. Familiarize yourself with the ports before insertion. Check the integrity of the balloons before you insert the tube.
• Place an endoscope protection mouthguard in place (to prevent biting of the tube). Cover the end of the tube with lubricating jelly, and, with the patient in the left semi-prone position, push the tube down, asking the patient to swallow (if conscious). If the tube curls up in the mouth, try again.
• Estimate the length of the tube to be inserted by measuring from the bridge of the nose to the earlobe and adding the distance from the nose to the xiphoid process. This should equate to at least 50–60cm; make sure that the tube is not coiled up in the back of the mouth.
• Inflate the gastric balloon with 250mL of water. Clamp the balloon channel. Then gently pull back on the tube until the gastric balloon abuts the gastro-oesophageal junction (resistance felt), and then pull further until the patient is beginning to be tugged by pulling. Note the position at the edge of the mouthpiece (mark with a pen), and attach with a sticking plaster to the side of the face.
• Tip: if the above fails, place the tube through the mouthguard to the back of the throat, and then follow with an endoscope. The endoscope will push the tube down the oesophagus and can be retroverted to directly visualize the gastric balloon being filled, before being removed.
• In general, the oesophageal balloon should never be used. Virtually all bleeding varices occur at the oesophagogastric junction and are controlled using the gastric balloon.
• Do not leave the balloon inflated for >12h, since this increases the risk of oesophageal ulceration.
• Obtain a CXR to check the position of the tube.
• The gastric channel should be aspirated continuously.
Patients should be warned of the risk of bleeding, pneumothorax, gall bladder puncture, failed biopsy, and shoulder tip pain, which may last several hours. The mortality is ~1:10 000.
• PT >3s prolonged.
• Platelet count <80 × 109/L or bleeding diathesis.
• Liver cancer (risk of tumour seeding).
Premedicate the patient with analgesia (e.g. 30–60mg of dihydrocodeine) before the procedure. The patient lies supine, with their right hand behind their head. Always carry out the liver biopsy under US guidance, especially if the liver is small and cirrhotic. The skin is cleaned, local anaesthetic infiltrated down to the liver capsule, and a liver biopsy needle is inserted when the breath is held in expiration. The biopsy itself takes about 5–10s and may cause shoulder tip pain.
A plugged biopsy can be performed when the PT is up to 6s prolonged, with a platelet count of >40 000mm3. The biopsy is done through a sheath, and the tract embolized using Gelfoam® to prevent bleeding.
Transjugular liver biopsy
This liver biopsy is taken through the hepatic vein, with secondary bleeding occurring into the circulation. It is not without risk, as the hepatic capsule may be punctured, leading to bleeding. It is used for patients in whom a prolonged PT or low platelet count precludes a normal liver biopsy.
A large introducer is placed into the IJV. A catheter is introduced through this and manipulated into the hepatic vein. The catheter is removed, leaving a guidewire in situ. A metal transjugular biopsy needle is passed over the wire and advanced into the hepatic vein. One has to avoid being too peripheral (risk of capsular puncture). The wire is removed, and the needle advanced while suction is applied. A biopsy is obtained by the ‘Menghini’ technique. The biopsies obtained are smaller and more fragmented than those obtained by conventional techniques.
• Uncontrolled bleeding of oesophageal or gastric varices.
• Diuretic-resistant ascites.
• Hepatic hydrothorax.
To decrease the portal pressure acutely, a shunt is placed between the hepatic vein and a portal vein tributary. Blood then flows from the high-pressure portal system to the lower-pressure hepatic venous system which drains into the IVC.
It is carried out in specialist centres and is technically quite difficult. It does not require a general anaesthetic, and it does not hinder future liver transplantation.
The IJV is catheterized, and a cannula passed through the RA into the IVC and into the hepatic vein. The portal vein is localized by USS, and a metal transjugular biopsy needle pushed through the liver from the hepatic vein into a portal vein tributary (usually the right portal vein). A wire is then passed into the portal vein and the metal needle withdrawn, leaving the wire joining the hepatic vein and portal vein. An expandable stent is then passed over the wire. A typical stent size is 8–12mm.
• Immediate mortality is 2–3%, usually from a capsular puncture and bleeding. The 4- to 6-week mortality in patients treated by TIPS for uncontrolled haemorrhage is up to 50% (from cirrhosis).
• Hepatic encephalopathy occurs in 20%.
• Failure to reduce portal pressure may occur if there are large extrahepatic shunts. These may need to be embolized.
Rarely used but does not require vascular access or anticoagulation. A clearance rate of ~10mL/min may be achieved.
• A peritoneal dialysis (PD) catheter (inserted under local anaesthesia).
• An intact peritoneal cavity free of infection, herniae, and adhesions.
• Features of peritonitis are: cloudy PD bag (99%), abdominal pain (95%), and abdominal tenderness (80%).
• Other features include: fever (33%), nausea and vomiting (30%), leucocytosis (25%), and diarrhoea or constipation (15%).
• Investigations: PD effluent cell count (peritonitis if >100 neutrophils/mm3); culture PD fluid (inoculate a blood culture bottle); Gram-stain PD fluid; FBC (for leucocytosis); blood cultures.
• All patients require antibiotics but may not require admission. The antibiotics used depend on Gram stain and culture results. A typical protocol would be ciprofloxacin or vancomycin, plus metronidazole. Patients with high fever with leucocytosis and/or who are systemically unwell warrant IV antibiotics.
• Gram –ve infection, in particular Pseudomonas, is associated with more severe infection.
• Peritonitis can lead to the development of an ileus.
• Patients may lose up to 25g of protein/day in severe cases and should receive adequate nutritional support.
• If the infection is resistant to treatment, consider removal of the Tenckhoff catheter and atypical organisms (e.g. fungi).
• Consider an underlying GI pathology, especially if multibacterial, Gram –ve organisms, or other symptoms.
Mild cases of fluid overload may respond to hypertonic exchanges (6.36% or 4.25% glucose), fluid restriction (1L/day), and large doses of diuretics (e.g. furosemide 500mg bd).
Other problems include poor exchanges, malposition of the catheter, omental blocking, fibrin deposition, and hyperglycaemia.
A blood flow of 250–300mL/min is needed across the dialysis membrane and leads to a clearance of 20mL/min.
• Vascular access: vascular access may be obtained using an arteriovenous shunt involving the radial artery or, more commonly, by using a Vascath which uses venous, rather than arterial, blood.
• Anticoagulation: heparin is normally used. If contraindicated, e.g. recent haemorrhage, then epoprostenol may be used but may cause hypotension and abdominal cramps.
• Haemodynamic stability: patients with MOF commonly develop hypotension during haemodialysis. This may be ameliorated by high-sodium dialysate and priming the circuit with 4.5% human albumin solution.
Complications of haemodialysis
Usually occurs within the first 15min of commencing dialysis. It probably involves activation of circulating inflammatory cells by the membrane, osmotic shifts, and possibly loss of fluid. Treatment: cautious fluid replacement and inotropes (watch for pulmonary oedema if overtransfused).
This occurs during the initial dialysis, especially in patients with marked uraemia, and is more common in patients with pre-existing neurological disease. Clinical features: headache, nausea and vomiting, fits, cerebral oedema. Treatment: treat cerebral oedema as described under Intracerebral haemorrhage, pp. [link]–[link]. Short and slow initial dialyses may prevent this.
This is caused by an IgE or complement response against the ethylene oxide component (sterilizing agent) or the cellulose component. Use of ‘biocompatible’ membranes, e.g. polysulfone, polyacrylonitrile (PAN), or dialysers sterilized by steam or gamma-irradiation may prevent further reactions.
Continuous arteriovenous haemofiltration (CAVH) implies bulk solute transport across a membrane and replacement. Continuous arteriovenous haemodiafiltration (CAVHD) involves pumping of dialysate across the other side of the membrane. For both, arterial blood (driven by arterial pressure) is continuously filtered at a relatively low flow rate (50–100mL/min). Continuous venovenous haemofiltration (CVVH) and continuous venovenous haemodiafiltration (CVVHD) involve pumping blood from a venous access to the dialysis membrane (150–200mL/min). The equivalent GFR obtained by these are 15–30mL/min. These are used most commonly on ITU. Both of these methods cause less haemodynamic instability and are particularly useful in patients with MOF.
A therapy directed towards removal of circulating high-molecular-weight compounds not removed by dialysis. Particularly used in the removal of antibodies or lipoproteins.
• Myasthenia gravis.
• Goodpasture’s syndrome.
• Severe hyperlipidaemia
• Multisystem vasculitis.
• Hyperviscosity syndrome (e.g. Waldenström’s macroglobulinaemia).
• HLA antibody removal.
Requires central venous access with a large-bore, dual-lumen cannula. Usually five treatment sessions are given on consecutive days. Plasma is removed and replaced with typically 2U of FFP and 3L of 4.5% albumin. IV Ca2+ (10mL of 10% calcium gluconate) should be given with the FFP. Febrile reactions may occur, as with other blood products. Plasmapheresis has no effect on the underlying rate of antibody production but is a useful treatment in acute situations such as Goodpasture’s syndrome and myasthenia gravis.
• For HUS and TTP, one must use FFP only (preferably cryodepleted), usually a minimum of 3L/day ( Thrombotic thrombocytopenic purpura and haemolytic uraemic syndrome, p. [link]).
• For hyperviscosity syndrome, a centrifugation system is required, rather than a plasma filter ( Hyperviscosity syndrome, p. [link]).
• For lipopheresis, there may be severe reactions if the patient is on an ACEI.
• An alternative to plasmapheresis is immunoabsorption, in which two columns are used in parallel. This may be used in the removal of HLA antibodies, anti-GBM disease, or multisystem vasculitis.
Biopsy is now performed using real-time US guidance by trained doctors (see Box 15.6).
• Bleeding diathesis—unless correctable prior to biopsy.
• Solitary functioning kidney.
• Uncontrolled hypertension, i.e. DBP >100mmHg.
• Urinary tract obstruction.
• Small kidneys, since it is unlikely to be helpful.
Prior to biopsy
• Check Hb, clotting screen, G&S serum.
• Ensure IVU or USS has been carried out to determine the presence and size of the two kidneys.
• Consent the patient; >1% risk of bleeding requiring transfusion.
• The biopsy is taken with the patient prone on the bed, with pillows under the abdomen. The lower pole of either kidney is visualized by US. A trucut biopsy is taken from the lower renal pole under sterile conditions with local anaesthesia. The biopsy is taken with the patient holding their breath at the end of inspiration (displaces the kidney inferiorly). Following biopsy, they should have bed rest for 24h to minimize the risk of bleeding, and the BP and pulse monitored half-hourly for 2h, 1-hourly for 4h, then 4-hourly for 18h.
• Send renal biopsy tissue for light microscopy, immunofluorescence, EM, and special stains (e.g. Congo red).
• Bleeding: microscopic haematuria is usual; macroscopic haematuria in 5–10%; bleeding requiring transfusion in 1%.
• Formation of an intrarenal arteriovenous fistula may occur.
• Severe loin pain suggests bleeding.
• Pneumothorax and ileus are rare.
pHi determination (gastric tonometer)
Patients in shock have reduced splanchnic perfusion and O2 delivery. The resulting mucosal ischaemia may be difficult to diagnose clinically until it presents as GI bleeding or the sepsis syndrome. The earliest change detectable following an ischaemic insult to the gut is a fall in intramucosal pH. Gastric mucosal pH parallels the changes in pH in other portions of the GIT, and monitoring this allows detection of gut ischaemia early.
A tonometer is essentially an NG tube with a second lumen leading to a balloon which lies within the mucosal folds of the stomach. The balloon is inflated with 0.9% saline for 30–90min. This allows CO2 from the mucosa to diffuse into the saline and equilibrate. The saline is then removed and analysed for pCO2, with simultaneous arterial blood [bicarbonate] measurement. pHi is then calculated using a modification of the Henderson–Hasselbalch equation.
Many synovial joints can be safely aspirated by an experienced operator. Knee effusions are common, and aseptic aspiration can be safely performed in the emergency department. The risk of inducing septic arthritis is <1 in 10 000 aspirations, but certain rules should be followed:
• Anatomical landmarks are identified.
• The skin is cleaned with alcohol or iodine.
• Local anaesthetic is applied to the area.
• A no-touch technique is essential.
• Aspirate the joint to dryness, by leaving the needle in the joint space and changing syringes.
Indications for synovial fluid aspiration in casualty
• Suspected septic arthritis.
• Suspected crystal arthritis.
• Suspected haemarthrosis.
• Relief of symptoms by removal of effusion in degenerative arthritis.
Contraindications to joint aspiration
• Overlying sepsis—never insert a needle through an area of cellulitis, as there is a risk of introducing infection into the joint.
• Bleeding diathesis.
• Prosthetic joints—must be aspirated in theatre by the orthopaedic surgeons
The patient lies with the knee slightly flexed and supported. The joint space behind the patella either medially or laterally is palpated, the skin cleaned, and a needle (18G, green) inserted horizontally between the patella and the femur using a no-touch technique. There is a slight resistance, as the needle goes through the synovial membrane. Aspirate on the syringe until fluid is obtained. (See Fig. 15.15a, b.)
Flex the elbow to 90°, and pass the needle between the proximal head of the radius (locate by rotating the patient’s hand) and the lateral epicondyle; or the needle can be passed posteriorly between the lateral epicondyle and the olecranon. (See Fig. 15.15c.)
Plantarflex the foot slightly; palpate the joint margin between the extensor hallucis longus (lateral) and the tibialis anterior (medial) tendons just above the tip of the medial malleolus. (See Fig. 15.15d.)
When synovial fluid is obtained:
• Note the colour and assess the viscosity.
• Microscopy for cell count and crystals (see Table 15.3).
• Gram stain and culture.
• AFB for suspected TB (although synovial fluid is usually non-diagnostic, and a synovial biopsy should be performed).
Table 15.3 Synovial fluid analysis
Leucocyte count (per mm3)
1000 (<50% PMN)
1–50 000 PMN
10–20 000 (poly-/mononuclear)
5–50 000 PMN
10–100 000 PMN
• Cerebral trauma (GCS score ≤8, compression of the basal cistern on CT, midline shift of >0.5mm on CT, non-surgical raised ICP).
• Acute liver failure (grade 4 coma with signs of raised ICP).
• Metabolic diseases with raised ICP (e.g. Reye’s syndrome).
• Post-operative oedema (after neurosurgery).
• After an intracranial haemorrhage (SAH or intracerebral).
ICP monitoring should be started before secondary brain injury in patients who are at risk of sudden rises in ICP and where it would influence management of the patient. These patients may be effectively managed in district hospitals.
• Uncorrectable coagulopathy.
• Local infection near the placement site or meningitis.
• There are several types of devices available (subdural, extradural, parenchymal, or intraventricular); parenchymal and intraventricular monitors carry a high risk.
• There are prepackaged kits available (e.g. the Codman® subdural bolt). This monitor is inserted in the prefrontal region, and the kit contains the necessary screws for creating a burr-hole, spinal needles to perforate the dura, etc.
• The ICP waveform obtained is a dynamic recording that looks similar to a pulse waveform, and is due to pulsations of the cerebral blood vessels within the confined space of the cranium, together with the effects of respiration.
• Cerebral perfusion pressure = MAP – ICP.
• The normal resting mean ICP measured in a supine patient is <10mmHg (<1.3kPa).
• The level which requires treatment depends, to some extent, on the disease—in benign intracranial hypertension, values of >40mmHg may not be associated with neurological symptoms; but in patients with cerebral trauma, treatment should be initiated when the ICP is >25mmHg.
• There are several types of pressure waves described, of which the most significant are ‘A waves’—sustained increases of the ICP lasting 10–20min up to 50–100mmHg (6–13kPa). These are associated with a poor prognosis.
• Readings of the ICP monitors should always be accompanied by careful neurological examination.
You will need
• Spinal needles and a manometer for measuring the opening CSF pressure.
• Dressing pack (gauze, drapes, antiseptic, gloves, plaster).
• Local anaesthetic (e.g. 2% lidocaine), three sterile bottles for collecting CSF, and a glucose bottle.
(See Fig. 15.16.)
Give antibiotics first if suspected meningitis ( Acute bacterial meningitis: assessment, p. [link]).
• Explain the procedure to the patient.
• Position the patient. This is crucial to success. Lie the patient on their left side if you are right-handed or on their right side if you are left-handed, with their back on the edge of the bed, fully flexed (knees to chin), with a folded pillow between their legs, keeping the back perpendicular to the bed. Flexion separates the interspaces.
• The safest site for LP is the L4–L5 interspace (the spinal cord ends at L1–L2). An imaginary line drawn between the iliac crests intersects the spine at the L4 process or L4–L5 space exactly. Mark the L4–5 intervertebral space.
• Clean the skin, and place the sterile drapes over the patient.
• Inject and anaesthetize the deep structures with 2% lidocaine.
• Insert the spinal needle (stilette in place) in the midline, aiming slightly cranially (towards the umbilicus), horizontal to the bed. Do not advance the needle without the stylet in place.
• You will feel the resistance of the spinal ligaments, and then the dura, followed by a ‘give’ as the needle enters the subarachnoid space. Replace the stylet before advancing.
• Measure the CSF pressure with the manometer and 3-way tap. Normal opening pressure is 7–20cmCSF. CSF pressure is with anxiety, SAH, infection, SOL, benign intracranial hypertension (BIH), and CCF.
• Collect 0.5–1.5mL of fluid in three serially numbered bottles, including a glucose bottle.
• Send specimens promptly for microscopy, culture, protein, glucose (with a simultaneous plasma sample for comparison), and, where appropriate, virology, syphilis serology, cytology for malignancy, AFB, oligoclonal bands (MS), CrAg, India ink stains, and fungal culture.
• Remove the needle, and place a plaster over the site.
• The patient should lie flat for at least 6h and have hourly neurological observations and BP measurements.
Complications of lumbar puncture
• Headache: common (up to 25%). Typically present when the patient is upright and better when supine. May last for days. Thought to be due to CSF depletion from a persistent leak from the LP site. Prevented by using finer spinal needles; keep the patient supine for 6–12h post-LP, and encourage fluids. Treat with analgesia, fluids, and reassurance.
• Trauma to nerve roots: rarer, but seen if the needle does not stay in the midline. The patient experiences sharp pains or paraesthesiae down the leg. Withdraw the needle, and if the symptoms persist, stop the procedure and seek expert help.
• Bleeding: minor bleeding may occur with a ‘traumatic tap’ when a small spinal vein is nicked. The CSF appears bloody (see CSF analysis, p. [link]), but the bleeding stops spontaneously and does not require specific therapy. Coagulopathy, severe liver disease, or thrombocytopenia carries the risk of subarachnoid/subdural bleeding and paralysis.
• Coning: herniation of cerebellar tonsils with compression of the medulla is very rare, unless the patient has raised ICP. Always get a CT brain scan prior to LP, and review this yourself if possible. Mortality is high, but the patient may respond to standard measures for treating this ( Raised intracranial pressure, pp. [link]–[link]).
• Infection: rare if a proper sterile technique used.
(See Table 15.4.)
Table 15.4 CSF analysis
Main cell type
• Normal values:
• Lymphocytes <4/mm3; polymorphs 0/mm3.
• Protein <0.4g/L.
• Glucose >2.2mmol/L (or >70% plasma glucose).
• Opening pressure <20cmCSF.
• A bloody tap is indicated by progressively fewer red cells in successive bottles and no yellowing of CSF (xanthochromia). The true WCC may be estimated by:
• True CSF WCC = CSF WCC – (blood WCC × CSF RBC)/blood RBC
• If the patient’s blood count is normal, subtract ~1 WBC for every 1000 RBCs). To estimate the true protein level, subtract 10mg/L for every 1000 RBCs/mm3 (be sure to do the count and protein estimation on the same bottle).
• SAH: ( Subarachnoid haemorrhage: assessment, p. [link]) xanthochromia (yellowing of CSF); red cells in equal numbers in all bottles. The RBCs will excite an inflammatory response (increasing CSF WCC), most marked after 48h.
Occupational exposures to bloodborne viruses (BBVs) in health-care workers can be divided into two groups: percutaneous (needle-stick) and mucocutaneous (e.g. through broken skin or via splashes into the eyes). High-risk body fluids include: blood, pleural fluid, peritoneal fluid, pericardial fluid, synovial fluid, amniotic fluid, human breast milk, CSF, saliva (in dentistry), semen, vaginal secretions, and unfixed tissues and organs (vomit, faeces, and urine only when contaminated with blood).
The major pathogens associated with needle-stick injuries and mucocutaneous exposures are:
Occupational exposures to BBVs can be caused by certain work practices such as:
• Not properly disposing of used needles.
• Recapping needles.
• Not using protective equipments, e.g. eye protection.
Assume that every patient is potentially infected with a bloodborne infection. The same precautions should be taken for every patient and every procedure.
• Cover skin cuts and abrasions with waterproof dressings.
• Never recap needles or pass sharps hand to hand.
• Always dispose of used needles promptly in sharps disposal containers at the point of use.
• Never leave sharps to be cleared up by others.
• Use eye protection. Ordinary spectacles offer inadequate protection. Use safety glasses, which fit over spectacles.
• Double-gloving reduces the risk of BBV transmission from a sharp injury.
• Use safer sharp devices; according to Health and Safety (Sharp Instruments in Healthcare) Regulations 2013: ‘the employer must substitute traditional, unprotected sharps with a “Safer Sharp” where it reasonably practicable to do so’.
Management of exposure incidents
• If the mouth or eyes are involved, wash thoroughly with water.
• If the skin is punctured, let the wound bleed and wash it with soap and water.
• Report to the occupational health department to arrange immediate assessment or, if out of hours, attend the A&E department.
Assessment of the risk of bloodborne virus transmission
Estimated seroconversion risks are:
• HBV: 30% for percutaneous exposure of a non-immune individual to HBsAg- and HBeAg-positive source.
• HCV: 1.9% for percutaneous exposure to HCV-infected blood with detectable HCV RNA.
• HIV: 0.3% for percutaneous exposure to HIV-infected blood.
• Percutaneous injury is higher risk than mucous membrane or broken skin exposure.
• Injury with a device directly from a source patient’s artery/vein.
• Injury from hollow-bore and wide-gauge needles.
• Deep injury.
• Visible blood on the device.
• High HIV viral load, or HBeAg in the source patient.
• Staff member inadequately immunized against hepatitis B.
Approaching source patients for bloodborne virus testing
• Due to the sensitivity of the issue, the source patient should not be approached by the exposed member of staff.
• Occupational health (or A&E if out of hours) will arrange this test.
Post-exposure prophylaxis for HIV
(See Box 8.4.)
• Risk assessment is carried out by occupational health (A&E if out of hours).
• PEP should be initiated as soon as possible—ideally within an hour, and generally within 72h of exposure, and continued for 28 days.
• Follow-up is carried out by occupational health, in accordance with local policy.
PEP for HBV
PEP for hepatitis B following significant occupational exposure depends on if the recipient has been immunized against hepatitis B and if adequate immunity has been achieved. Risk assessment is carried out by occupational health (A&E if out of hours).