• Users Online: 104
  • Print this page
  • Email this page


 
 Table of Contents  
REVIEW ARTICLE
Year : 2019  |  Volume : 57  |  Issue : 4  |  Page : 299-307

Maintenance and Management of an ophthalmic operation theater in sub-Saharan Nigeria


1 Department of Vitreo Retina, Aravind Eye Hospital, Chennai, Tamil Nadu, India
2 Former Director, Eye Project, Tulsi Chnarai Foundation, Abuja, Nigeria
3 Department of Cornea, Aravind Eye Hospital, Chennai, Tamil Nadu, India
4 Former CEO, Tulsi Chanrai Foundation, Abuja, Nigeria

Date of Submission09-Oct-2019
Date of Acceptance17-Oct-2019
Date of Web Publication26-Dec-2019

Correspondence Address:
Dr. Prabu Baskaran
Department Vitreo Retina, Aravind Eye Hospital, Poonamallee High Road, Chennai - 600 007, Tamil Nadu
India
Login to access the Email id

Source of Support: None, Conflict of Interest: None


DOI: 10.4103/tjosr.tjosr_84_19

Get Permissions

  Abstract 


In today's era, the operation theatre (OT) is like the heart of an ophthalmic hospital set up. The maintenance and management of this sensitive area is quite often missed or not taught during the training period. It becomes imperative to have a sound knowledge about OT maintenance and management, particularly if the set up is located in a remote area with limited resources and all the more it makes sense if an ophthalmologist has to manage the entire set up single handedly. The objective of this article is to meet this requirement. Based on accepted clinical guidelines and our own experience, we have to compiled most of the important aspects of OT set up and maintenance.

Keywords: Nigeria, operation theater management, sterilization


How to cite this article:
Baskaran P, Rangarajan A, Ramakrishnan S, Gadok SS. Maintenance and Management of an ophthalmic operation theater in sub-Saharan Nigeria. TNOA J Ophthalmic Sci Res 2019;57:299-307

How to cite this URL:
Baskaran P, Rangarajan A, Ramakrishnan S, Gadok SS. Maintenance and Management of an ophthalmic operation theater in sub-Saharan Nigeria. TNOA J Ophthalmic Sci Res [serial online] 2019 [cited 2020 Jan 19];57:299-307. Available from: http://www.tnoajosr.com/text.asp?2019/57/4/299/273986




  Introduction Top


In the current scenario, the operation theatrer (OT) has become the vital point of any ophthalmic hospital set up. The proper management of this sensitive area generally requires a team effort. It becomes imperative to have a sound knowledge about OT maintenance and management, particularly if the setup is located in a developing country with limited resources. The objective of this article is to meet this requirement. We ran a dedicated eye hospital in Kebbi, a remote sub-Saharan area in northern Nigeria, and cater mainly to the elimination of cataract blindness in this part of the world. Our center performs 3000–3500 cataract operations with intra-ocular lens implantation, in a year. Based on accepted clinical guidelines and our own experience in this region, we have tried to compile certain important tenets of OT setup and maintenance, as described under ten headings as follows.

Operation theater layout

An efficient OT begins right from the planning and construction of the OT complex. The OT complex should be located well away from the roadside so as to avoid dust and noise. The operating room should be in the center of the OT complex.

Modern-day OT incorporates zoning of areas within the OT complex [Figure 1]:
Figure 1: Layout of the operation theatre complex

Click here to view


  • Outer zone – reception area which is accessible to all persons and supplies
  • Clean zone – space for the OT staff to move around after changing into an OT suit. It should include changing room, space for shifting of patients and block room
  • Aseptic zone – consists of scrub-up area, sterilization room, and operating room, the latter being the most sterile area
  • Disposal zone – area for disposal of bio-hazardous waste.


The autoclave room should be away from the main operating room, so that moist heat generated from the autoclave does not enter the operating room. It has to be noted that this moisture is a vital factor for the growth of microorganisms.

Physical set up

Do's

  • Glaze tiling (larger tiles are preferred so as to reduce the number of joints between the tiles)
  • Epoxy painting of walls (contains antibacterial properties)
  • Corners should preferably be rounded so as to prevent the accumulation of dust and facilitate easy cleaning
  • Doors should be airtight
  • Separate entries to the operating room for the patients, staff, sterile items, and exit for waste disposal.


Don'ts

  • No false ceilings – this is ideal, but in Africa and other developing nations where most constructions are with false ceilings, we suggest the use of plastic washable sheets which are easier to maintain and clean instead of the usual thermocol or plaster of Paris
  • No openable windows
  • No wooden materials
  • No ceiling fans.


Physical parameters

Ventilation

Twenty–twenty-five air exchanges per hour are necessary, out of which at least five should be fresh air from outside.[1],[2],[3] Split air conditioner (AC) or central AC is mandatory for this purpose. Laminar airflow or positive pressure ventilation is the best options, but their availability in a routine set up is restricted by their cost. Air purifiers such as high-efficiency particulate air filters can be made available. Adding ultraviolet (UV) light further helps in purifying the air. Since UV light is toxic to human beings, it is switched on only after the last person leaves the OT and switched off before the first person enters the OT. Temperature was set at 18°C–24°C.[1],[3] Relative humidity was set at 55%–80%.[1],[2] Bacteria-containing particles of air within 30 cm of the operation site should not exceed 10/cubic meter, and should not be more than 20/cubic meter in the rest of the OT.[1]

Block room requirements

  • The patient should be given local anesthesia in the lying-down (supine) position only
  • Emergency tray with all basic necessary medicines should always be available. It should contain multiple boxes segregated and named alphabetically depending on their contents. This makes handling easy during an emergency [Figure 2]
  • An oxygen cylinder should always be available. Its functionality should be checked every day, and it is preferable to have an additional cylinder as a spare in the block room
  • An intravenous (IV) stand should be erected near the blocking table; a normal saline bottle is always kept attached to it
  • Items like IV catheter, oxygen mask, etc., should be readily available
  • The presence of a general physician is mandatory.


By following all these tenets meticulously, one can reduce the reaction time of the medical team when an emergency hits.
Figure 2: Emergency drugs arranged in alphabetical order

Click here to view


Instrument cleaning

Surgical instruments should be cleaned as soon as possible after their use. The quality of water used for cleaning is very important. The initial rinse may be normal piped drinking water.[4] However, the water for the final rinse should have a higher quality with a minimum of particles and minerals dissolved in it. Although the preferred method for water treatment is distillation, it is difficult to obtain large quantities of water by this method. A single distillation table-top unit can be used for obtaining distilled water at least for the final steps of cleaning. Other methods, such as water softening, de-ionizing, and reverse osmosis (RO), are used in cleaning and sterilization units to obtain the required quantity of high-quality water.[4]

After their use, sharp and blunt instruments should be first separated. They are then cleaned with either piped drinking water or water purified by RO and made available as running water through taps. Running water allows the removal of debris and remnant tissues from the instruments.

All the instruments are then soaked in a large bowl containing a mixture of antiseptic solution (e.g., Dettol) and distilled water in 1:40 ratio for 10 min. The active ingredient in Dettol is chloroxylenol. Fine tipped instruments are cleaned with a soft brush like a clean toothbrush [Figure 3].
Figure 3: Cleaning instruments with a soft brush in dettol solution

Click here to view


Next, the instruments are transferred to an ultrasonic cleaner. Antiseptic solution like Dettol should be added along with distilled water (ratio of 1:40) in its chamber so as to avoid cross-contamination between the instruments. The water height should be few inches above the height of the tray being kept in the chamber [Figure 4]. An ultrasonic cleaner liberates high-frequency sound waves at the rate of 20,000/s, generating bubbles and vacuum zones. The resulting action is called cavitation. It thoroughly cleans every part of the instrument. Instruments should undergo cleaning in this manner for 15 min. If this apparatus is not available, the four-bowl technique can be followed. Instruments are passed through four consecutive bowls, the first bowl containing a combination of Dettol and water as specified above, followed by three bowls containing distilled water.
Figure 4: Ultrasonic cleaner

Click here to view


The automated instrument rinsing system is ideal for cleaning cannulas [Figure 5]. If it is not available, cannulas have to be manually flushed three times with distilled water and three times with air, alternately. Manual flushing is best done with larger syringes like 20 ml syringe.
Figure 5: Automated instrument rinsing system for cannulated instruments

Click here to view


Finally, the instruments should be taken out and kept on a table [Figure 6] for drying with ambient or preferably hot air.[5] Hairdryers can be used for this purpose.[6] The cloth should not be used for drying instruments as it can damage the sharp edges of instruments and leave behind unwanted lint fibers. All detachable parts must be disassembled and kept for drying. Drying of instruments before autoclaving is extremely important since steam cannot thoroughly percolate the lumen of the cannulas if they are wet. The presence of water droplets makes subsequent sterilization incomplete and ineffective. In addition, remnant water droplets can themselves be a source of microorganisms [Table 1].
Figure 6: Instruments segregated and kept for drying

Click here to view
Table 1: Classification of chemical indicators[13],[14]

Click here to view


After drying, each instrument has to ideally be checked for its working condition under well-illuminated magnascopes [Figure 7]. Sharp tips of instruments should be capped with bits of an infant feeding tube or IV tubings and kept in an instrument roll. The instrument roll is a piece of linen stitched such that it has small individual pouches for each and every instrument [Figure 8]. This roll is securely tied with a piece of string attached to it.[6] These rolls are placed in surgical autoclave bins, which are loaded into the autoclave. This specialized bin has a latch which controls the apertures in its body. Just before the bins are loaded into the autoclave, this latch is opened, exposing these apertures, thus helping in effective steam percolation. Immediately after the bins are removed from the autoclave chamber following sterilization, this latch should be closed [Figure 9] and [Figure 10].
Figure 7: Magnascope

Click here to view
Figure 8: Instrument roll with individual pouches being packed for sterilization

Click here to view
Figure 9: Surgical autoclave bin with latch opened: apertures exposed

Click here to view
Figure 10: Surgical autoclave bin with latch closed: apertures sealed

Click here to view


Instrument sterilization

Autoclave

An autoclave or steam sterilizer works under the principle of steam under pressure. There are two main types of conventional steam sterilizers:

  1. Gravity displacement type – holding time of 30 min at 121°C at 15 pounds per square inch (PSI) pressure[7],[8] [Figure 11]
  2. High-speed prevacuum type – holding time of 4 min at 132°C at 30 PSI pressure[7],[8] [Figure 12].
Figure 11: Gravity displacement type steam sterilizer

Click here to view
Figure 12: High-speed prevacuum type steam sterilizer

Click here to view


Pressure settings may vary slightly depending on the sterilizer used. When possible, follow manufacturers' recommendations. Furthermore, at constant temperatures, sterilization times vary depending on the type of item (e.g., metal vs. rubber vs. plastic, items with lumens, etc.), whether the item is wrapped or unwrapped, and the type of sterilizer.[7],[9]

The autoclave water indicator should be monitored before starting the process. Water is added up to the zero mark only [Figure 13]; excess water will result in wetting of surgical gowns, drapes, etc.
Figure 13: Autoclave water indicator to be filled up to zero mark only

Click here to view


Note that the autoclave cycle will restart if there is any electric failure during the cycle or even if there is a short transition of power source from standard electrical supply to generator.

Once the cycle is complete, as indicated by an audible sound, the pressure valve should be opened, so that the pressure inside the autoclave chamber equalizes atmospheric pressure. The exhaust fan should be used to avoid condensation of vapors inside the sterilization room. Alternatively, if facilities are available, the steam vent can be connected to a channel that can release the steam into the outside atmosphere.

The autoclave lid can then be opened. The bins, after removal from the autoclave, should be kept on wire shelves that allow the free flow of air around them so that they cool without condensation.

Once the bins are cooled, they should be wrapped in a polythene bag and stored in a dry, dust-protected place, above the waist level. The bins should not be piled on each other.[6] All steam-sterilized items should be used within 48 h of sterilization.[9],[10]

Flash sterilization is a modification of conventional steam sterilization (either gravity or prevacuum type) in which the flashed item is placed in an open tray to allow rapid penetration of steam. By original definition, flash sterilization is sterilization of an unwrapped object at 132°C for 3 min at 27–28 lbs of pressure.[7] This type of sterilization is considered acceptable for processing items that cannot be packaged, sterilized, and stored before use.[7] It is also used when there is insufficient time to sterilize an item by the preferred package method and for sterilization of unwrapped surgical instruments for emergency purposes (e.g., dropped instruments, etc.).[8] It has to be noted that flash is not recommended as a technique for routine sterilization of surgical instruments.[11],[12]

Monitoring of autoclave

A separate logbook for the autoclave should be maintained, documenting the date and time of the start and end of every cycle, the items being sterilized, and the color change noted on the indicator. The operating surgeon should verify and sign the autoclave register before starting the surgery.

Biological indicators are the most ideal tools for sterilization monitoring, but using biological indicators for each package would be very expensive and the results would be available only after about 24 h. Due to this, monitoring of every cycle is done with chemical indicators as a routine. The ANSI/AAMI/ISO 11140-1: 2005/(R) 2010 document defines six classes of chemical indicators.[13],[14] The classification has no hierarchical significance.

Bowie Dick test[10] (Class 2 indicators) should be done in an empty load, at the start of every sterilization day, to ensure that there is no airlock inside the autoclave chamber [Figure 14]. Commonly used autoclave tapes are Class 1 indicators or process indicators that indicate if a determined package went through the process; however, it does not guarantee that the package has been sterilized. Color change occurs by mere exposure to the sterilization process and not by the satisfaction of set parameters [Figure 15]. These indicators are used externally on all packages.
Figure 14: Bowie-Dick test sheets before exposure (blue color) and after exposure (uniform black color) indicating no airlock within the autoclave chamber

Click here to view
Figure 15: Autoclave tape before exposure (white diagonal stripes on the tape) and after exposure (black diagonal stripes on the tape)

Click here to view


Class 5 integrating or Class 6 emulating indicators should be used for further strengthening the reliability of sterilization because these are internal indicators which change color only when all the parameters, namely pressure, temperature, and time get satisfied. Currently, Class 5 integrating indicators are the most accurate of the chemical indicator classes [Figure 16] when compared to the performance of biological indicators.[14]
Figure 16: Class 5 indicators before (left) and after (right) completion of the autoclave cycle. Note, purple color of 3 arrows and central oval has fully changed to green color

Click here to view


Biological monitoring[1],[7],[10] using Geobacillus stearothermophilus spores for steam sterilizers [Figure 17] is recommended to be done on a weekly basis.[15]
Figure 17: Bioindicator for steam sterilization - Geobacillus stearothermophilus spores

Click here to view


We currently use a combination of Class 1, 2, and 5 chemical indicators and biological indicators at our own center.

Ethylene trioxide sterilization

Ethylene trioxide (ETO) is a toxic gas and a very effective sterilizing agent. It can be used for sterilizing all heat sensitive articles. Objects should be dry before ETO; otherwise, a toxic layer of ethylene glycol will form on their surface. Adequate aeration time after running the cycle is necessary to allow dissipation of free toxic gas. The parameters are as follows:[7]

  1. Gas concentration – 450–1200 mg/l
  2. Temperature – 37°C–63°C
  3. Relative humidity – 40%–80%
  4. Cycle time – 1–6 h


  5. Aeration time – Depends on technique (mechanical aeration for 8–12 h at 50°C–60°C or ambient

  6. room aeration for 7 days at 20°C)
  7. Shelf life – 1 year.


Biological monitoring using Bacillus subtilis[1],[10] or Bacillus atrophaeus[7] for ethylene oxide sterilization is recommended once weekly.

Operation theater cleaning

Fumigation

The standard agent used for fumigation is formaldehyde. Routinely, 500 ml of 40% formalin solution is mixed with 1000 ml of distilled water for every 1000 cubic feet.[1] An aerosol machine or a fogger is used for fumigation as they produce microparticles. A fogger would be the better option because the particle size generated by it is smaller. The timer should be set for 30 min. After fumigation is complete, the room should be sealed airtight with adhesive tape for 24 h. Fumigation is done once a week or more frequently, as judged appropriate by the surgeon if the surgical load at a given center is high. Apart from routine weekly fumigation, it is considered mandatory to perform fumigation following any septic surgery, new construction, or reconstruction of OT or if culture report reveals any pathogenic microorganisms in the OT.

Glutaraldehyde and formaldehyde combination can also be used[10] as a change, once in 2 months, to avoid resistance developing to one particular agent. Liquid ammonia solution (1 L ammonium solution plus 1 L of water for every liter of 40% formaldehyde used[2]) can be used to absorb the formalin fumes when the OT is opened after fumigation.

Although fumigation has been described as a time-tested method of OT disinfection; it is presently being considered obsolete in many countries due to the toxic nature of formalin. Symptoms of excess exposure include respiratory irritation, burning and watering of eyes, itchy, runny, or stuffy nose, dry or sore throat, and headache. Contact with formaldehyde can cause skin irritation and dermatitis. Formaldehyde is also listed by the International Agency for Research on Cancer as carcinogenic to humans.[16],[17] Newer chemicals (Bacillocid Rasant, Virkon etc.) are now being introduced as less hazardous and quicker methods of disinfection but they are associated with financial constraints. Active ingredients in Bacillocid rasant include Glutaral 100 mg/g, benzyl-C12-18-alkyldimethylammonium chloride 60 mg/g, didecyl-dimethylammonium chloride 60 mg/g. Virkon is a multi-purpose disinfectant. It contains oxone (potassium peroxymonosulphate), sodium dodecylbenzenesulfonate, sulphamic acid, and inorganic buffers.

Rather than fumigation, more stress is now being laid on cleaning of the OT floor with antiseptic solution, especially in the vicinity of the immediate surgical field. Antiseptic, such as Savlon, a combination of cetrimide and chlorhexidine, can be used. This is done not only at the end of the operation day but also in between surgeries at regular intervals. Ideally, this may be done after each operation, though this may not be practically possible in centers with a large volume of surgeries.

Daily cleaning

The operating room is always cleaned first, followed by the scrub-up area, sterilization area, block room, and the common corridor. The operating microscope head should be cleaned with Bacillocid solution (a combination of chemically bound formaldehyde, glutaraldehyde, benzalkonium chloride, and alkyl urea derivatives). Lenses are kept covered after cleaning with lens cleaning solution (available as a combination of ammonium hydroxide, denatured ethyl alcohol, sodium lauryl sulfate, and water). Operating tables, chairs, trolleys, IV stands, etc., are cleaned with antiseptic solution. Door handles or push plates require special attention due to higher likelihood of contamination. OT walls (up to six feet height from the floor) and the wall around the scrub sink[18] also require special attention.

The floor of the OT is always mopped last, using a two-bucket technique. One bucket is filled with antiseptic solution (Dettol: water – 1:40 ratio) and the second one with warm water. The mop is first dipped in the antiseptic solution, and the floor is mopped, then it is dipped in the warm water, and the procedure is repeated.

The UV light is switched on when the last person leaves the OT and is switched off before the first person enters the OT on the next day. This is to ensure an aseptic environment even when no activity is going on.

Weekly cleaning

In addition to daily cleaning routines, cleaning of the ceiling as well as any wall-mounted items should be done on a weekly basis. All the articles are removed from their shelves and cleaned. Filters of the ACs are removed, cleaned with detergent solution, and dried well before fixing back.[10]

Monthly cleaning

All the equipment and cupboards are taken out from the OT and cleaned well. The walls and floor of the OT are thoroughly mopped with antiseptic solution.

Water tank cleaning

It is preferable to have a water tank or reservoir that is dedicated exclusively to the OT. This tank is cleaned once in 15 days.[10] First, the water has to be drained completely, followed by cleaning of the base and sides of the tank with a detergent solution. It is then allowed to dry under sunlight for half an hour. The detergent is then washed with water and cleaned once again with antiseptic solution, further allowing it to dry for another half an hour. Finally, the tank is rinsed well with water, twice, before filling the tank with clean water.

The water filter has to be cleaned every week with detergent solution, and it has to be changed periodically.

Microbiological evaluation

Multiple culture swabs are taken every week from areas such as microscope head, head end of OT tables, trolleys, AC vents, OT wall, OT floor, IV stands, scrub sink, sterilization room wall and floor, autoclave chambers, block room table, water used for scrubbing, air from the operating room, etc., along with one control swab. The control swab can be contaminated or sterile (this will be known to the sterilization in-charge only), and this will be decided on a random basis. The microbiologist should not be aware of the swab site or the nature of the control swab. He/she should only report against the swab number. This ensures the reliability of the reports.

Operation theatre waste disposal

The OT staff should be aware of color coding of waste disposal bags, especially while dealing with bio-hazardous waste. Incineration has been advocated as an effective and cheap method of hospital waste disposal.[19] Recently, the attention has been directed towards preventing air pollution from incineration, and towards finding alternative technologies for the treatment of medical waste. Infective waste may be autoclaved prior to disposal.[20] Other viable techniques include gasification, heat disinfection of certain clinical wastes prior to disposal in a landfill, etc.[21] Sharp items like needles, blades, etc., should be placed intact into puncture-resistant containers, without being manipulated by hand, to avoid needle-stick injuries.[20]

Surgical precautions

  • The operating surgeon and assisting staff should wear rubber boots or covered shoes[22]
  • The surgeon should wear a plastic or rubber sheet[22] or waterproof surgical apron inside the gown so that he/she does not come in contact with body fluids
  • The surgeon should wear protective spectacles[22] even if he is an emmetrope to prevent exposure of his/her eyes to patients' body fluids.


Patient care

Case sheets of patients requiring special attention, such as hypertensive, diabetic, cardiac, or one-eyed patients, should be marked with colored stamps or labels for ease of identification at every level and to alert the surgeon, especially in a volume surgical setup. Patients are made to wear clean OT gowns, caps, and foot covers. Before entering the OT complex, it is desirable to have the patients clean their feet with antiseptic solution [Figure 18]. Preoperative cleaning of the eye to be operated and the periocular skin with 5% povidone-iodine solution is done once in the waiting area [Figure 19] and repeated before block. After blocking, the patient is kept in the lying down posture for 15 min before being taken into the operating room.
Figure 18: The patient cleaning feet with antiseptic solution before entering operation theater complex

Click here to view
Figure 19: Cleaning with povidone-iodine in the waiting area of the operation theater complex

Click here to view


Inside the operating room, after painting the immediate surgical area once again with povidone-iodine, the eye to be operated is draped with a sterile eye towel and a sterile plastic disposable adhesive eye drape is applied. An aperture for the eye is cut open in this drape, and the cut edges are securely tucked under the eyelid margins, thus, effectively isolating the eyelashes and lid margins from the surgical area. This eliminates the need for trimming the eyelashes before surgery. In his book “Eye Surgery in Hot Climates,” author J. Sanford-Smith writes, “A much cheaper alternative is to autoclave a piece of plastic cellophane 'clingfilm' spread out on paper. This can be placed across the eyelids and lashes and a small hole cut in the middle for the speculum. After using the drapes should be washed in soapy water and sun-dried before packing and sterilizing.” Such alternative methods may be considered in the presence of financial or material limitations. During the functioning of the OT, especially in a high volume surgical setup, it is to be noted that consumables should not be shared between two trolleys or two patients.

After the surgery, the patient should be taken to the recovery room and kept on the bed for 15 min. Then, he/she is taken to the ward in a wheelchair. These measures reduce the chance of postural hypotension.

In this article, we have tried to provide certain basic practical points for day-to-day functioning of an ophthalmic OT while adhering to accepted guidelines. These caveats can be followed by eye care professionals and OT personnel, even in setups with limited resources and in remote areas. This will ensure high-quality ophthalmic care to all our patients.

Declaration of patient consent

The authors certify that they have obtained all appropriate patient consent forms. In the form the patient(s) has/have given his/her/their consent for his/her/their images and other clinical information to be reported in the journal. The patients understand that their names and initials will not be published and due efforts will be made to conceal their identity, but anonymity cannot be guaranteed.

Financial support and sponsorship

Nil.

Conflicts of interest

There are no conflicts of interest.



 
  References Top

1.
Guidelines For: Quality Cataract Management in Secondary Level Eye Centres. Developed by Venu Eye Institute & Research Centre and Sewa Rural along with Sightsavers International. Available from: http://www.sightsavers.org/in_depth/quality_and_learning/learning/13172_Guidelines%20for%20quality%20cataract%20management.pdf. [Last accessed on 2012 Jan 18].  Back to cited text no. 1
    
2.
Ram J, Kaushik S, Brar GS, Taneja N, Gupta A. Prevention of postoperative infections in ophthalmic surgery. Indian J Ophthalmol 2001;49:59-69.  Back to cited text no. 2
[PUBMED]  [Full text]  
3.
Srinivasan V, Thulasiraj RD. Ophthalmic instruments and equipments. In: A Hand Book on Care and Maintenance. 2nd ed. Madurai city: Sri Aurobindo Ashram Press, Aravind Eye Care System; 2003. p. 40.  Back to cited text no. 3
    
4.
Cox I, Stevens S. Care of ophthalmic surgical instruments. Community Eye Health 2000;13:40-1.  Back to cited text no. 4
    
5.
Rutala WA, Weber DJ. The Healthcare Infection Control Practices Advisory Committee (HICPAC). Guideline for Disinfection and Sterilization in Healthcare Facilities. Centers for Disease Control and Prevention; 2008. p. 58-63, 111-2. Available from: http://www.cdc.gov/hicpac/pdf/guidelines/Disinfection_Nov_2008.pdf. [Last accessed on 2012 Jan 19].  Back to cited text no. 5
    
6.
Tietjen L, Bossemeyer D, McIntosh N. Infection Prevention Guidelines for Healthcare Facilities with Limited Resources. JHPIEGO, an Affiliate of Johns Hopkins University; 2010. p. 11-2. Available from: http://www.reproline.jhu.edu/english/4morerh/4ip/IP_manual/ipmanual.htm. [Last accessed on 2012 Feb 02].  Back to cited text no. 6
    
7.
Standardized Clinical Protocols; Sterilization Protocol. Developed by LAICO, Aravind Eye Care System. Available from: http://v2020eresource.org/DisplayDetails.aspx?bid=M00000000001120001. [Last accessed on 2012 Feb 04].  Back to cited text no. 7
    
8.
Steam Chemical Indicator Classifications 3MTM Sterile U Tutorials. Available from: http://multimedia. 3m.com/mws/mediawebserver?mwsId=66666UuZjcFSLXTtmXTVOXM6EVu QEcuZgVs6EVs6E666666--&fn=70-2009-7519-4.pdf. [Last accessed on 2012 Feb 02].  Back to cited text no. 8
    
9.
Living Organisms for Sterility Assurance-Biological Indicators. 3M Sterile U Tutorials. Available from: http://multimedia. 3m.com/mws/mediawebserver?mwsId=SSSSSu7zK1fslxtUn82BMY_Uev7qe17zHvTSevTSeSSSSSS-- and fn=70-2010-7243-9.pdf. [Last accessed on 2012 Feb 15].  Back to cited text no. 9
    
10.
Basu RN. Issues involved in hospital waste management – An experience from a large teaching institution. J Acad Hosp Adm 1995;7-8:79-83.  Back to cited text no. 10
    
11.
Prajna L, Charu Priya R, Noor S. Sterilization in Ophthalmic Practice. Vision 2020 e Resource. Available from: http://laico.org/v2020resource/files/equip_sterilise.pdf. [Last accessed on 2012 Jan 20].  Back to cited text no. 11
    
12.
Phillips G. Microbiological aspects of clinical waste. J Hosp Infect 1999;41:1-6.  Back to cited text no. 12
    
13.
Stevens S. Control of infection in ophthalmic practice. Community Eye Health J 2003;16:40-1.  Back to cited text no. 13
    
14.
Ram J. Reducing cataract-related complications. Indian J Ophthalmol 1999;47:153-4.  Back to cited text no. 14
[PUBMED]  [Full text]  
15.
ANSI/AAMI/ISO 11140-1: 2005/(R) 2010 Sterilization of Health Care Products – Chemical indicators – Part 1: General Requirements. 2nd ed. Available from: http://www.aami.org. [Last accessed on 2012 Mar 08].  Back to cited text no. 15
    
16.
The American Society of Cataract and Refractive Surgery Ad Hoc Task Force on Cleaning and Sterilization of Intraocular Instruments. Recommended Practices for Cleaning and Sterilizing Intraocular Surgical Instruments. Supplement to EyeWorld; March, 2007. Available from: http://www.eyeworld.org/ewsupplementarticle.php?id=200. [Last accessed on 2012 Mar 10].  Back to cited text no. 16
    
17.
Basics of Sterilization-Tutorial. Published Online by Case Medical Inc. Available from: http://www.casemed.com/caseacademy/downloads/CASDF003.pdf. [Last accessed on 2012 Mar 09].  Back to cited text no. 17
    
18.
Huys J. Cleaning of Equipment and Materials to be Sterilized. The Sterile Supply Cycle: Cleaning. Published online by World Forum for Hospital Sterile Supply. Available from: http://www.wfhss.com/html/educ/sbasics/sbasics0102_en.htm. [Last accessed on 2012 Mar 10].  Back to cited text no. 18
    
19.
Sandford-Smith J. Operating theatre procedures and equipment. In: Eye Surgery in Hot Climates. 3rd ed. F A Thorpe; 2004. p. 44. Available from: http://www.cehjournal.org/files/eshc/eysurhc_ch3.pdf. [Last accessed on 2012 Feb 07].  Back to cited text no. 19
    
20.
Operative Operation Theatre. Manual Developed by SEWA Rural, Jhagadia; 2006. Available from: http://www.gujhealth.gov.in/Images/got%20dr.%20kalpanaben%20eng%20new.pdf. [Last accessed on 2012 Mar 10].  Back to cited text no. 20
    
21.
International Agency for Research on Cancer. IARC Monographs on the Evaluation of Carcinogenic Risks to Humans. Volume 88 (2006): Formaldehyde, 2-Butoxyethanol and 1-tert-Butoxypropan-2-ol. International Agency for Research on Cancer; June, 2004. Available From: http://monographs.iarc.fr/ENG/Monographs/vol88/index.php. [Last accessed on 2012 Mar 11].  Back to cited text no. 21
    
22.
Formaldehyde and Cancer Risk. National Cancer Institute at National Institutes of Health. Available from: http://www.cancer.gov/cancertopics/factsheet/Risk/formaldehyde. [Last accessed on 2012 Mar 11].  Back to cited text no. 22
    


    Figures

  [Figure 1], [Figure 2], [Figure 3], [Figure 4], [Figure 5], [Figure 6], [Figure 7], [Figure 8], [Figure 9], [Figure 10], [Figure 11], [Figure 12], [Figure 13], [Figure 14], [Figure 15], [Figure 16], [Figure 17], [Figure 18], [Figure 19]
 
 
    Tables

  [Table 1]



 

Top
 
 
  Search
 
Similar in PUBMED
   Search Pubmed for
   Search in Google Scholar for
 Related articles
Access Statistics
Email Alert *
Add to My List *
* Registration required (free)

 
  In this article
Abstract
Introduction
References
Article Figures
Article Tables

 Article Access Statistics
    Viewed27    
    Printed0    
    Emailed0    
    PDF Downloaded3    
    Comments [Add]    

Recommend this journal


[TAG2]
[TAG3]
[TAG4]