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Year : 2001  |  Volume : 49  |  Issue : 1  |  Page : 59-69

Prevention of postoperative infections in ophthalmic surgery

Department of Ophthalmology Post Graduate Institute of Medical Education and Research, Chandigarh, India

Correspondence Address:
Amod Gupta
Department of Ophthalmology, Post Graduate Institute of Medical Education and Research, Chandigarh - 160 012, India

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Source of Support: None, Conflict of Interest: None

PMID: 15887720

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Postoperative endophthalmitis is a serious, vision-threatening complication of intraocular surgery. Better instrumentation, surgical techniques, prophylactic antibiotics and better understanding of asepsis have significantly reduced the incidence of this complication. Postoperative endophthalmitis may occur as an isolated event or as a cluster infection. Topical antibiotics, preoperative periocular preparation with povidone-iodine combined with a sterile operating room protocol significantly reduce the incidence of isolated postoperative endophthalmitis. The role of antibiotics in the irrigating fluid and subconjunctival antibiotics remains controversial. Cluster infections on the other hand are more likely to occur due to the use of contaminated fluids/viscoelastics or a breach in operating room asepsis. Prevention of postoperative endophthalmitis requires strict adherence to operating room norms, with all involved personnel discharging their assigned roles faithfully.

Keywords: Eye surgery, asepsis, cluster infection, isolated infection, operating room, postoperative endophthalmitis

How to cite this article:
Ram J, Kaushik S, Brar GS, Taneja N, Gupta A. Prevention of postoperative infections in ophthalmic surgery. Indian J Ophthalmol 2001;49:59-69

How to cite this URL:
Ram J, Kaushik S, Brar GS, Taneja N, Gupta A. Prevention of postoperative infections in ophthalmic surgery. Indian J Ophthalmol [serial online] 2001 [cited 2023 Dec 6];49:59-69. Available from: https://journals.lww.com/ijo/pages/default.aspx/text.asp?2001/49/1/59/22669

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Maintenance of asepsis is imperative to ensure safe surgery and to minimize postoperative infection and its disastrous consequences. Postoperative endophthalmitis remains one of the most devastating complications of eye surgery, leading to a marked loss of vision in over 80% of affected cases.[1] The incidence appears to be influenced by geographical location, the aseptic technique used and preoperative antibiotic prophylaxis.

Prior to 1950, the average incidence of endophthalmitis was approximately 1%[2]. Based on a review of several reports, Allen and Mangiaracine found the incidence of postoperative infection following cataract extraction to be as high as 11.6/1000.[3] From 1950-64, the use of preoperative topical antibiotics brought down the incidence to 2.1/2000 cases, as seen at the Massachusetts Eye and Ear Infirmary.[3] The most current figures on the incidence of culture-proven endophthalmitis are available in a review of over 30,000 procedures done at the Bascom Palmer Eye Institute, Miami, where it was found to be 0.9/1000 cases.[4] This improvement is generally attributed to better instrumentation, a better understanding of aseptic techniques and the prophylactic use of antibiotics.

Postoperative endophthalmitis may occur clinically as an isolated event or as a cluster of infections in the form of a surgical epidemic. [5, 6] The most common infecting organism is Staphylococcus epidermidis with a reported incidence in two studies of 70% and 33% respectively. [7, 8] However, in most cases, the source of the infecting organism cannot be conclusively identified.

Discussed below are factors affecting isolated and cluster post-operative infections under three major headings: operating room (OR) layout and disinfection, sterilisation of instruments, and sterile surgical protocol.

  Operating Room (OR) Layout and Disinfection Protocol Top

  Operating room layout Top

Modern OR design incorporates zoning of areas within the OR complex.[9] Important aspects of any OR layout include location, design, proper ventilation, and separation of the sterile zone from the non-sterile areas. An excellent review of OR layout has been provided by Sharma et al.[10] The OR should be planned such that it is away from inpatient areas and other sources of infection. Often, an upper floor in the building is considered appropriate. For strict asepsis, an eye OR should preferably not be shared with any other surgical discipline. Contamination from a hospital construction environment has been documented as the source of an Aspergillus endophthalmitis epidemic.[11]

The major zones of an OR complex are: a) Outer zone, b) Clean zone, c) Aseptic zone, and d) Disposal zone.

a)The outer zone is a reception area and is accessible to all persons and supplies.

b)The clean zone is the space for circulation of OR staff after changing. It encompasses the (i)changing room located near the entrance of the OR complex; and (ii)transfer zone which is the space where patients are shifted from the transfer point to the OR.

c)The aseptic zone is the sterile area within the OR complex consisting of the scrub and gowning area, the preparation room and the OR. The OR should have one entrance and a separate opening towards a sterile area marked for instrument packing and sterilisation. Floors and walls should preferably be of non-porous material with minimum joints to enable proper cleaning and carbolisation. The head end of the operating table should be directed away from the entrance. The space should be so designed that the ventilatory flow in this zone is directed outwards from the operating field.

d)The disposal zone is the area where used equipment and supplies are processed. Disposal of biohazardous waste is also done in this zone.

  Ventilation Top

Air decontamination is important. The OR should be well ventilated and the circulating air should preferably be filtered. High Efficiency Particulate Air (HEPA) systems remove most microorganisms ranging in size from 0.5-5.0μ.[12] The principle of ventilation in the OR is the delivery of positive pressure filtered air in a vertical unidirectional flow over the operating table. The current United States Public Health Service minimum requirements for optimum OR air is as follows: temperature between 18-24°C, humidity 55-80%, and 25 changes per hour.[12] Fridkins et al[13] reported four patients who contacted Acremonium kiliense endophthlmitis due to defective ventilation in the OR. Laminar airflow curtains or a radial exponential airflow pattern away from the operating field are especially helpful.

In the surgical OR providing facilities for most forms of surgery, the recommended bacterial count of air should not exceed l/ft3(35.5/m3).[14] Air entering the OR from filters should not contain more than 0.5 /m3 of bacteria-containing particles. Furthermore, the bacteria-containing particles of air within 30 cm of the operation site should not exceed 10/m3, and should not be more than 20 /m3 in the rest of the OR.[14]

  Cleaning and disinfection of the OR Top

Cleaning, disinfection and sterilisation are the cornerstones in ensuring OR asepsis. The terms are independent of each other and each needs to be understood separately.

a) Cleaning the OR

Cleaning essentially means the removal of foreign matter (e.g. soil, organic matter) from the concerned surface. The first consideration is cleaning the OR. All surfaces should be free from visible dirt. It is normally accomplished with water, mechanical scrubbing and detergents. This is often neglected in comparison to disinfection and sterilisation, but is of equal importance. Unless an article is mechanically clean, there will not be sufficient surface contact between it and the decontaminating agent, and sterilisation will not be accomplished. Cleaning also reduces the bacterial count, though it will not disinfect or sterilise.

b) Disinfection and sterilisation of the OR

Disinfection is a process of freeing the concerned object of all pathogenic microorganisms which may cause infection during its use. Sterilisation is a process that eliminates all living organisms from the treated object.[15] It is impractical to attempt to sterilise the entire OR and equipment, therefore current practices concentrate on disinfection. The measures commonly used are discussed below.

i) Phenolic compounds

Phenol (carbolic acid) is one of the oldest known germicides. The hundreds of compounds derived from it constitute phenolic compounds, which are good bactericides and are active against fungi.[15] They are sometimes virucidal but are not sporicidal, except at temperatures over 100°C. They are very stable and remain effective after mild heating and prolonged drying.

However, these agents are not recommended as high-level disinfectants due to their lack of activity against bacterial spores and lack of published efficacy data of available formulations. This class of compounds is used for decontamination of the OR and for noncritical medical and surgical items. The floor and 2 meters of OR walls should be mopped with phenolic solution. Similarly, wet mopping of all OR tables, mats, instrument trolleys, stools, chairs and supply shelves with phenol followed by a wipe down with 70% alcohol is an effective daily decontaminating regimen.[16] Anaesthetic equipment like endotracheal tubes, airways and suction apparatus should be disinfected after every use. This problem is circumvented these days by the use of disposable items.

b) Formaldehyde fumigation

Formaldehyde is an effective agent commonly used to sterilise the OR. The efficacy of the process is uncertain at temperatures below 20°C and relative humidity below 70%.[17] For optimum OR disinfection, formaldehyde fumigation is recommended fortnightly as a routine, and at the end of an operating session of a grossly infected case.

All apertures in the room should be sealed with adhesive tape prior to fumigation. The gas is liberated by spraying or heating formalin or solid paraformaldehyde. For each 1000 cubic feet of space (28.3m3), 500 ml of 40% formaldehyde in one litre of water is put into an electric boiler or a large bowl placed on a electric hot plate with safety cut-out when boiling dry. The OR is sealed after turning on the boiler or hot plate. After fumigation, the room is kept closed for at least 8-10 hours. Subsequently, ammonium solution is introduced and left in the room for a few hours to neutralise the formaldehyde (one litre ammonium solution plus one litre of water for every litre of 40% formaldehyde used.)

Other methods of fumigation are[18]

a) Permanganate method

Five ounces of potassium permanganate for every 1000 cu.ft. of space are placed in a jar and on top of this 10-15 ounces of 40% formalin diluted with an equal amount of water is poured. As soon as the reagents are mixed, a violet effervescence takes place and formaldehyde is set free.

b) Paraform method

On heating formalin, the aldehyde changes into the solid polymeride - paraform. Gas is generated by heating paraform tablets. 25-30 tablets are required for every 1000 cu. ft. of space.

c) Formalin spray / vapouriser

Aeromax vapouriser can be used to fumigate an OR. 250 cc of 40% formalin dissolved in 5000 cc tap water makes a dilution of 1:20. One litre of the solution is used per 1000 cu.ft. of space. This is vapourised over half an hour. Spraying is not a satisfactory substitute for vaporisation of formaldehyde by boiling as the fine aerosol has poor penetration.[17]

A recommended daily regimen for cleaning and disinfection of the OR is illustrated in [Table - 1].

A less often discussed issue is the problem of ensuring microscope sterility during ophthalmic microsurgery. Usually, sterilisable handle protectors, or some form of a plastic sterilisable drape for the entire microscope is used. A simple and effective alternative has been designed for the Zeiss® operating microscope.[16] A metal bracket mounts in the accessory shoe of the microscope, and a rigid plastic sterilisable portion is readily attached to the microscope from below with three sterilisable screws. The entire assembly can be attached very quickly (15 seconds). The system ensures full sterility in the operating field, provides a sterile surface for handling and moving the microscope by the surgeon or his assistant and at the same time allows open access to accessories like cameras and observer tubes.

  OR etiquette Top

A high standard of discipline is necessary for the safe conduct of any surgery. Personnel entering the OR complex should be strictly controlled. Anyone with overt infection should be barred. All persons entering the OR should change into freshly laundered clothing. Hair and beards should be clean and be well covered by caps and masks. High-filtration disposable masks are to be worn at all times when within the aseptic zone. Female health workers should take special care to trim nails and remove jewellery when working within the theatre complex. All persons must wash their hands thoroughly before entering the OR.

It is desirable to restrict all persons other than the staff from entering the OR. In the electronic age, it is better for students and other trainees to observe surgery from a remote location on closed circuit television rather than crowd around the surgeon's table. This is particularly advantageous when using the operating microscope with a camera mounted on a beam splitter.

  OR waste disposal Top

Operating room biohazardous waste including infected linen, disposable syringes and needles, intravenous (IV) drip sets, IV fluids, and infected and diseased excised pathological tissue poses a significant health hazard to the OR personnel and public. Safe disposal is imperative to prevent spread of infection and possible recycling of hazardous disposable products.

Over the last decade, the disposal of operating room and hospital waste has received much attention. Incineration has been advocated as a viable method of hospital waste disposal.[19] Recently, attention has been directed at preventing air pollution from incineration, and to find alternative medical waste treatment technologies. These options include gasification, steam sterilisation or heat disinfection of certain clinical wastes prior to disposal in a landfill.[20]

  Sterilisation of Instruments Top

Surgical instruments and drapes need to be sterilised adequately. It must be appreciated that sterilisation is an absolute term, and there is no such thing as partial sterility. The goal of instrument processing is to protect patients by preventing cross-contamination from instruments. The processing includes a sequence of steps to remove microbes on contaminated instruments and to keep them sterile for future use. The American Operating Room Nursing (AORN) Recommended Practices Committee[21] has provided guidelines for the care and cleaning of surgical instruments.

Instruments need thorough cleaning after every surgery before being sterilised. Fine microsurgical instruments are best cleaned by an ultrasonic cleaner. These contain liquids through which sound waves pass at a frequency of 100,000 Hz or more. [10, 22] These waves generate submicroscopic bubbles which then collapse, creating a negative pressure on the particles in the suspension. Bacteria disintegrate and protein matter is coagulated by this action.[18] It is especially useful for instruments with hinged areas or serrated edges which are difficult to clean by hand.

Despite their advantages, ultrasonic cleaners are not recommended for telescopes of endoscopes, or other lumened devices such as phaco or irrigation and aspiration handpieces. Such instruments require irrigation using a high-pressure water jet. It is important that all lumened instruments be flushed with cool water before being subjected to heat which would coagulate proteins and block the channel.

Sterilisation can be done by physical or chemical methods, of which the former is more reliable.

  Physical agents Top

  Sterilisation by heat Top

  Dry heat Top

Hot air by itself is an inefficient sterilising agent since it is a poor conductor and does not penetrate well. A temperature of 160°C for one hour or 180°C for 20 minutes will sterilise the contents by a destructive oxidation of cell constituents. [10, 18, 22] The one-hour holding period is timed after the temperature reaches 160°C. Its usefulness is limited to some sharp instruments, which would otherwise be damaged by moist heat.

  Autodaving Top

This method is more effective than dry heat and requires lower temperatures in a given time. Autoclaving at 121°C for 15 minutes at 15 psi pressure effectively kills most microorganism. A temperature of 134°C at 34 psi pressure sterilises instruments within 3 minutes.[23] Temperature-sensitive detectors must always be used to ensure adequate autoclaving. Autoclaving is suitable for sterilisation of most metallic ophthalmic instruments except sharp knives and fine scissors. Autoclaving irrigating solution bottles may kill heat-labile microorganisms but bacterial spores may still survive, since steam cannot penetrate the bottles.

  Sterilisation by filtration Top

This method is used to sterilise heat labile fluids. It employs a 0.22μ. on-line micropore filter for irrigating solutions, fluid-gas exchange, and intraocular air/gas injection. [24, 25] Microorganisms are retained in part by the small size of the filter pores and in part by the adsorption on the pore walls during the passage of the fluid through the filter. Their main limitation is failure to retain viruses which are usually much smaller than the pores of the filters currently available. Gills[25] has reported a reduction in the rate of endophthalmitis after cataract surgery from 1-2 per 1000 to 1 in 8000-10000 surgeries after filtration.

  Chemical agents Top

  Glutaraldehyde 2% (Cidex®) Top

This is an effective steriliser for instruments that cannot be autoclaved. It is non-corrosive, does not impair the sharpness of cutting instruments and may be used with plastic, aluminium and rubber. Glutaraldehyde in a 2% aqueous solution is more effective and less irritant. It is stable in acidic solution and even more so in alkaline solution. Therefore, the commercial preparation Cidex® is supplied with a separate alkaline buffer containing a rust inhibitor, which is added before its use. It is effective against vegetative pathogens in 10 minutes and resistant pathogenic spores in 3 hours.[23] It is very effective against the tubercle bacillus.

The low surface tension allows for easy penetration to inner surfaces and it can be readily removed by rinsing. Thorough rinsing of all sterilised material is mandatory because residual Glutaraldehyde is extremely irritating to tissues. Sterilisation of lumencontaining instruments such as irrigating cannulae by glutaraldehyde is not recommended. Recent studies indicate that such use may lead to corneal endothelial cell damage and uveitis. Courtright et al[26] have reported corneal oedema in 37% of 111 patients who underwent extracapsular cataract extraction. The corneal oedema was maximum in the first 48 hours after surgery. Multivariate analysis showed that the day of surgery was the only characteristic significantly associated with the presence of corneal oedema. A simulation of the disinfection technique used revealed significant levels of glutaraldehyde in instruments with small lumens even after thorough rinsing.

Glutaraldehyde 2% is suitable for sterilisation of sharp instruments, which cannot be autoclaved. These must then be rinsed serially two to three times in trays filled with Ringer's lactate or normal saline (0.9%), to eliminate the residual chemical from the instruments.

  Ethylene oxide (ETO). Top

ETO is widely used commercially to sterilise single-use items. It is also used in some large hospitals for resterilising packaged heat-sensitive devices like sharp knives and blades. Gas sterilisation using ethylene oxide is effective and safe for heat-labile tubings, vitrectomy cutters, cryoprobes, lightpipes, laser probes, diathermy leads and most disposable items for cost reduction. Sterilisation is done by a process known as alkylation in which a hydrogen atom is replaced by a hydroxyl ethyl radical within a protein molecule. When a sufficient number of positrons within molecules of microorganisms have been alkylated, death ensues. The process kills all microorganisms, including tubercle bacilli and spores.[27]

It penetrates very well and is relatively non-toxic compared to formalin. The easy penetrability allows the article to be prepackaged. Aeration is essential for all items sterilised by ETO. The duration varies and is dependent upon the absorbency of the load and the temperature and air exchange rate of the aeration facility. If higher aeration temperatures are used, for instance 55°C, twelve hours is usually sufficient, but periods of up to one week are usually necessary at room temperature. After aeration, the equipment can be stored sterile for extended periods of time.

To be effective and safe, several parameters must be considered. For effective sterilisation, the minimum concentration required is 400-1000mg/L (the concentration of ETO is measured in mg of gas per litre of space). Required exposure time can be decreased by increasing the temperature. Most automatic sterilisers employ temperatures of 40-60°C. Moisture enhances the diffusion of the ETO gas. Usually steam is introduced into the steriliser before admitting the ethylene oxide. Blood, pus and other proteinaceous materials act as barriers to ETO. Equipment must therefore be cleaned well before sterilisation.

The type of packaging used is important because ETO must be both capable of penetrating it and of being removed from it. Polyethene of 200-gauge thickness satisfactorily wraps equipment for long-term storage and its transparency allows one to see what is in the package.

The time required varies with the type of steriliser depending upon all the above factors, and also the type of load. Sterilisation time could vary from 1-12 hours.[27] These parameters must always be checked for compliance with the manufacturers' design specifications and initial performance tests.

A typical ETO sterilisation cycle includes the following steps:[28]

  • Labeling each article with the date and time and an indicator for sterilisation before loading

  • Air removal with a vacuum pump (70-cm Hg vacuum)

  • Heating to the required temperature, usually 45-55°C

  • Steam humidification of the load to a relative humidity of 60%

  • Exposure to the ETO for the prescribed period (5 psi for 12 hours or 10 psi for 6 hours.)

  • Gas removal by 70 psi vacuum.

  • Air flush by filtered air repeated 3-4 times to re-establish atmospheric pressure.

  • Aeration to elute residual ETO from the load.

Advantages of ethylene oxide are reliability, wide applicability, minimal damage to sterilised objects, and the feasibility of pre-packaging and long-term storage. Disadvantages include expense, the need for thorough training of personnel, and the need to adequately aerate equipment for extended periods of time before use.

  Acetone Top

Acetone is a potent bactericidal agent and is useful for routine disinfection of surfaces. This was once considered the method of choice for sterilising sharp instruments, but its effectivity is being increasingly questioned. Drews[29] has postulated that the poor results reported could be due to its relative ineffectiveness in the diluted form, and emphasized the need to use it in a concentrated solution. It has been found to be ineffective against fungi and spore-bearing bacteria and fungi even after 20 minutes.[30] It may be useful in situations when autoclaving or gas sterilisation is not feasible or available.

  Plasma sterilization Top

This new modality of sterilisation of instruments has been introduced recently for heat-sensitive medical devices. This process is claimed to have overcome many of the safety concerns associated with ETO sterilisation, and process times are much shorter.

Gas plasma is a highly excited body of gas produced by the application of energy to gas under vacuum. The ions and molecules within the plasma collide to produce free radicals, which are capable of interacting with microorganisms to disrupt their function. This principle has been used to develop a number of systems for sterilising heat sensitive instruments and other medical devices. The best known of these systems is the Sterrad (Advanced Sterilisation Products, California). This system used a low temperature (<50°C) hydrogen peroxide gas plasma. [31, 32]

The Sterrad process is initiated by creating a vacuum. A small volume of hydrogen peroxide in aqueous solution is injected from a cassette. This is vaporised and dispersed throughout the chamber and load. A further reduction of pressure is then applied and an electric field created using radio waves. This generates gas plasma from hydrogen peroxide. The free radicals so produced react with and destroy microorganisms present on pre-cleaned, dry accessible surfaces. The process is sporicidal, bactericidal, virucidal and fungicidal. It is applicable to both thermostable and thermolabile materials. Though not yet widely used in the Ophthalmic OR, its main advantage is that it can be used at temperatures below 50°C.The gas plasma process has advantages of low process times, viz. 50-80 min., so aeration is unnecessary, and no toxic emissions or residues are said to result from the process.

  Flash sterilization Top

Emergency sterilisation may be required occasionally if an instrument is dropped or unintentionally left out of the tray. Flash sterilisation is performed at 132°C at 28 1b of pressure for 3 minutes for metal instruments in a steriliser.[33] The recommended minimum exposure time for linen, rubber, plastic and lumen containing articles should be 10 minutes for gravity-displacement and 4 minutes for pre-vacuum cycles. However, the practice should be restricted to emergency situations only, since the margin of safety is lower. It is to be remembered that these parameters are the minimum for sterilisation, and any deviation from the prescribed exposure time, temperature or pressure can produce non-sterile items.

  Clean air storage hood. Top

A laminar flow clean air hood for the sterile storage of ophthalmic instruments in an operating room was evaluated in a Mayo Clinic study[34]. A total of 10,524 surgical procedures performed with the instruments stored in this hood were retrospectively analysed. The incidence of endophthalmitis in these cases was 0.076% (8 cases). No clusters of infection were identified. The use of such a laminar flow clean air hood provides access to surgical instruments in a high volume operating room without exposing patients to a high risk of endophthalmitis.

  Monitoring sterilisation protocols. Top

All sterilisation procedures must be monitored meticulously by appropriate means for optimum effectiveness.

The efficacy of a disinfectant or antiseptic may be measured by comparison with that of phenol under given conditions. The "phenol coefficient" is determined in parallel tests that compare killing times observed with suspensions of Salmonella typhi exposed to known concentrations of the test substance and killing times observed with suspensions exposed to known concentrations of phenol. This is the Rideal-Walker test. Organic matter in the form of dried yeast or dried human faeces may be included in the test system using the Chick-Martin test, [23, 28] to stimulate conditions under which disinfectants are expected to work.

The efficiency of a surface disinfectant is judged in terms of its ability to inactivate a known number of a standard strain of a pathogenic Staphylococcus on a given surface within a given time.

Monitoring sterilisation is more difficult. Sterilisation process indicators (e.g. temperature charts, pressure gauges) are used to indicate inadequate process conditions. Chemical and biological indicators are used to monitor the effectiveness of sterilisation. The biological indicators come closest to an ideal monitor because they integrate all of the sterilising parameters involved, such as time, temperature, pressure and packaging. Bacillus stearothermophilus, a thermophile that requires to be cultivated at 55°C to 60°C, is a suitable test organism; its spores are killed at 121°C in about 12 minutes. Chemical detectors such as Bowie-Dick tapes show a change of colour or shape after exposure to a sterilising temperature when applied to packs and articles in the load. The tape develops diagonal lines when exposed for the correct time to the sterilising temperature. [12, 23]

For ETO sterilisation, biological indicators comprising Bacillus subtilis var niger (NCTC 10073) spores dried onto suitable carriers and contained within the load, such as Line and Pickerill Helix®, are necessary for verification. Similar indicators are required for validation of sterilisation with the gas plasma steriliser.[28]

  Sterile Surgical Protocol Top

  Pre-operative assessment for patient related factors: Top

In isolated infections, patient-related factors predominate. Diabetic patients and those with blepharo-conjunctivitis, dry eyes or atopic disease are at a higher risk of postoperative infections as they have a higher rate of carriage of Staphylococcus aureus.[35] Other patient related preoperative risk factors include blepharitis, conjunctivitis, dacryocystitis, lacrimal drainage abnormalities, ocular surface disorders, host immuno-suppression and even upper respiratory tract infections in children.[36-38] The presence of unexplained fever or malaise must be thoroughly investigated for the possibility of an infective focus like a dental abscess, and must be adequately treated prior to surgery. It is also important to identify and treat any preexisting eye disease before surgery.

It is impossible to sterilise the external eye completely, and since the main source of infection is the patient's own ocular flora, it is imperative to try and reduce colony counts on the lids and conjunctiva preoperatively. The ocular surface and adnexa are the main sources of bacteria in culture-proven cases of endophthalmitis. Using microbial DNA analysis, Speaker and coworkers[39] have demonstrated that the main source of infection is the patient's own ocular flora. Gram-negative bacteria account for about 16% of cases, and fungi for a much smaller proportion of infections. Olson et al,[40] in their review of patients of postoperative infectious endophthalmitis, documented rates of 90%, 7% and 3% respectively for gram-positive aerobic bacteria, gram-negative bacteria and fungi. Anaerobic bacteria also inhabit the ocular surface and can cause endophthalmitis.[35] Thus it makes good clinical sense to reduce the microbial load in the immediate preoperative period.

The first step in achieving this objective is to emphasize the importance of a hot scrubbing bath with a head wash on the day of surgery. Thorough facial washing with a medicated soap helps to clean the surgical area prior to disinfection with chemical agents.

  Preoperative use of prophylactic antibiotics. Top

The role of prophylactic antibiotics administered both topically and subconjuctivally in reducing postoperative infection has been documented. [41, 42] Topical instillation of antibiotics in the conjunctival sac is a simple and minimally invasive method of delivering drugs to the anterior segment. Loading doses of topical antibiotics and frequent application (every 15-30 minutes) have been found to provide prolonged therapeutic levels in the cornea and aqueous humour.[43] Topical antibiotics should begin 24 hours before surgery and used 6-8 times during the day. Instillation of topical antibiotics for more than 24 hours may lead to replacement of the patients' own flora by more virulent microorganisms.

  Preoperative preparation and the role of povidone-iodine. Top

Speaker and Menikoff,[44] in a significant breakthrough showed that a single topical application of 5% povidone-iodine solution reduced the incidence of endophthalmitis by a factor of three. Povidone-iodine is bactericidal in 30 seconds and need not be irrigated out of the eye before surgery. [44, 45] This has become a routine practice with most ophthalmic surgeons during intraocular surgery.

Although cilia-trimming was once considered helpful in reducing postoperative infection, the present trend is not to trim cilia for intraocular surgery. This has followed the widespread practice of isolating lashes with sterile adhesive drapes. Cleaning the lids, lid margins and adjacent skin with povidone-iodine 5% is an effective method of eliminating microbes.[46]

  Scrubbing and the use of gloves Top

It is important to scrub adequately and use a nailbrush. One should scrub one's hands and arms till above the elbows. It takes 7-8 minutes to scrub with soap alone. A hand disinfection system using chlorhexidine reduces the rate of noso-comial infections more effectively than using alcohol and soap.[47] With povidone-iodine or chlorhexidine solution, scrubbing twice for 1-2 minutes each is adequate. The proper method of wearing gown and gloves is also important. Use of sterile gloves helps to reduce the transfer of organisms from the surgeons hands, skin and nails. After wearing sterile gloves, it is important to wash hands with Balanced Salt solution (BSS) or Ringer's lactate to remove talc from the gloves. Inflammatory anterior chamber reaction secondary to contaminated talc and adsorbable dusting powder has been reported.[48] One should also keep in mind perforations in the gloves which are found in a significant number.[49]

  Sterile drapes to isolate eyelashes/lid margins Top

Use of a sterile drape to cover the face, ocular adnexa and isolate eyelashes and lid margins helps to reduce the passage of microorganisms into the eye.[46] After painting the lid and surrounding skin with povidone-iodine 5% twice and allowing it to dry, the adhesive drape is applied after widely opening the lids. The drape is cut between the lids and the eye speculum applied in such a way that the drape is under the speculum arms, which isolate the cilia.

  Surgical procedure Top

Many factors are implicated in the occurrence of endophthalmitis including the patient's own immunity and the quality of surgery. Prolonged surgery (longer than 60 minutes), use of prolene haptic intraocular lenses, inadvertent ocular penetration during extraocular surgery and vitreous loss have been specifically documented to be risk factors for the development of endophthalmitis.[50] An intact posterior capsule usually acts as a barrier and prevents infection. The risk of developing infection is high if the integrity of the posterior capsule is lost, since it allows easy access for microorganisms into the vitreous cavity.[46] Vitreous loss requires introduction of additional instruments into the eye, which contributes to the increased risk of infection. All precautions should be undertaken to prevent a posterior capsule rupture.

Postoperative risk factors include poor wound construction or closure leading to a wound leak, iris prolapse or vitreous incarceration in the wound, exposed sutures, suture removal under inadequate aseptic conditions and the presence of a thin filtering bleb.[51]

  Irrigating fluids and viscoelastics Top

The fluids for intraocular and intravenous use such as BSS, Ringer's lactate, etc. should be inspected for intact packing and for any obvious bacterial or fungal contamination. Any visible particulate matter should render a bottle unsafe for use even if its sterile packing seems undisturbed. A similar evaluation of containers of viscoelastic material such as methylcellulose 2% or sodium hyaluronate is mandatory before use. Several reports of cluster postoperative endophthalmitis have been reported implicating contaminated intraocular solutions. [5, 6, 11,]

A cluster infection is defined as the occurrence of two or more than two infections at a time, or the occurrence of repeated postoperative infection. Contaminated fluids,[52-54] tubings, intraocular lenses,[5] and viscoelastics[6] are known to have caused cluster infections. These infections usually occur as a result of a breach in standard OR protocol. Isolated reports of cluster infections are available tracing the source in the ventilation system causing Acremonium endophthalmitis.[13] Pseudomonas aeruginosa endophthalmitis has been reported from contaminated automated surgical equipment such as the internal pathways of a phaco-emulsifier.[55]

In cluster infections, microbiological surveillance teams culture specimens for bacteria or fungi taken from environmental sites, the OR floor and walls, water source, OR personnel, irrigating fluids, viscoelastics, intraocular lenses, surgical equipment and the autoclaving equipment. Usually, despite a thorough search, the source of infection remains elusive.

Even with a sterile surgical technique, infection may occur from many other sources. It has been shown that bacteria may be recovered from culture of anterior chamber aspirates at the end of cataract surgery in up to 43% of cases even in the presence of preoperative antibiotic prophylaxis and aseptic techniques.[56] Lacunae in our knowledge include lack of information regarding how many bacteria are required to cause endophthalmitis and how low the bacterial counts have to be in order to ensure a negligible risk of postoperative infection.

  Prophylactic antibiotics during surgery Top

The role of antibiotics in irrigating solutions is controversial. [57, 58] The antibiotic protocol is usually aimed at providing protection against Gram-positive bacteria, which are the organisms most often cultured in endophthalmitis cases (75-90% of positive cultures).[59] Vancomycin 10 mg in a 500 ml bottle is the recommended dosage.[60] However, there are reports that this practice makes no difference to the incidence of postoperative endophthalmitis.[61] Another danger sounded by the US Centers for Disease Control and Prevention is the emergence of vancomycin-resistant coagulase-negative Staphylococcus and Enterococcus strains.[62]

Other considerations are the cost implications and the risk of human error during constitution of the required solution. This is especially true when using aminoglycosides, as inadvertent injection of toxic doses could result in macular infarction and endothelial toxicity. Moreover, it is very difficult to evaluate treatment or prophylactic measures for very low incidence complications such as endophthalmitis.

Subconjunctival injection of antibiotics at the end of surgery helps to reduce postoperative infection particularly in the setting of the developing world. A single injection of high doses of most antibiotics maintains therapeutic aqueous levels for 4-6 hours.[63] The definite efficacy is still questioned.[64] A 10-year retrospective review of culture-proven, acute-onset postoperative endophthalmitis by Aaberg et al[65] showed that 23 of these cases had received prophylactic subconjunctival antibiotics at the conclusion of surgery. Fourteen cases were sensitive to the antibiotics used and 9 were resistant.

  Surgery of infected cases Top

All infected cases should be operated in a separate OR, despite some recommendations to the contrary.[12] The tubings, instruments and sheets used for infected cases must be cleaned thoroughly and sterilised adequately before reuse. An added concern is the need for sterilisation or high level disinfection of medical devices contaminated with blood from patients infected with HIV or HBV, or with respiratory secretions from a patient with pulmonary tuberculosis. For this purpose, freshly activated glutaraldehyde 2%, 70% ethanol or isopropyl alcohol, 6% hydrogen peroxide, 4% formalin, 2% povidone-iodine or 1000 ppm of free chlorine have been recommended.[66] Thus, all semicritical medical equipment should be treated as if contaminated with blood-borne pathogens, and should be processed by the same rigorous technique.

It is evident that ensuring eye surgery free from the catastrophe of intraocular infection involves a whole team, each member of which must perform his/her assigned role faithfully. All staff working in the OR should be well versed with sterilisation norms and techniques being followed in their own theatre. Abiding by a few rules and ensuring OR discipline goes a long way in providing safe ocular surgery to patients.

  References Top

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  [Table - 1], [Table - 2]

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