1. Cleaning glassware and reusable syringes and needles
Instructions for cleaning:
— glass containers (Erlenmeyer flasks, beakers, test-tubes)
— pipettes
— microscope slides
— coverslips
— reusable syringes and needles.
Glass containers
New glassware
Glassware that has never been used may be slightly alkaline. In order to neutralize it:
● Prepare a bowl containing 3 litres of water and 60 ml of concentrated hydrochlo- ric acid (i.e. a 2% solution of acid).
● Leave the new glassware completely immersed in this solution for 24 hours.
● Rinse twice with ordinary water and once with demineralized water.
● Dry.
Dirty glassware
Preliminary rinsing
Rinse twice in cold or lukewarm water (never rinse bloodstained tubes in hot water).
If the glassware has been used for fluids containing protein, it should be rinsed immediately and then washed (never allow it to dry before rinsing).
Soaking in detergent solution
Prepare a bowl of water mixed with washing powder or liquid detergent. Put the rinsed glassware in the bowl and brush the inside of the containers with a test-tube brush (Fig. 3.57). Leave to soak for 2–3 hours.
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| Fig. 3.57 Cleaning dirty glassware |
Rinsing
Remove the articles one by one. Rinse each one thoroughly under the tap, then soak them all in a bowl of ordinary water for 30 minutes.
Rinse each article in a stream of clean water. (Do not forget that traces of detergent left on glassware can lead to false laboratory results.)
Draining
Place containers (beakers, flasks, measuring cylinders) on the pegs of a draining rack. Place test-tubes upside-down in a wire basket.
Drying
Place the glassware in wire baskets and dry in a hot-air oven at 60 °C. Alternatively, place the baskets in a sunny spot in the laboratory and cover them with a fine cloth.
Plugging
The clean dry glassware should be put away in a cupboard to protect it from dust. It is recommended that glass containers be plugged with non-absorbent cotton wool or their mouths covered with small caps made from newspaper (Fig. 3.58) or, preferably, thin sheets of paraffin wax or clinging plastic, if available.
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| Fig. 3.58 Plug or cover glassware to protect it from dust |
Pipettes
Immediate rinsing
Once a pipette has been used, rinse it immediately in a stream of cold water to remove blood, urine, serum, reagents, etc.
Soaking in water
After rinsing, place the pipettes in a large, plastic measuring cylinder (or bowl) full of water. If the pipettes have been used to measure infected material, leave them in a cylinder full of disinfectant solution (e.g. a quaternary ammonium compound or 1% bleach solution; see pages 84 and 85) for 4 hours.
Soaking in detergent and rinsing
Follow the instr uctions given above for soaking and rinsing of laborator y glassware.
Blocked pipettes
1. Put blocked pipettes in a cylinder filled with dichromate cleaning solution (reagent no. 20). Slide them carefully into the solution and leave for 24 hours.
2. The next day, pour the dichromate solution into another cylinder (it can be used four times).
3. Hold the cylinder containing the pipettes under the tap and rinse thoroughly.
4. Remove the pipettes one at a time. Check that the obstruction has been washed away. Rinse again.
5. Leave to soak in ordinary water for 30 minutes, then change the water and soak for a further 30 minutes.
Warning: Dichromate cleaning solution is highly corrosive and should be used with extreme care. If it is accidentally splashed on the skin or clothing or into the eye(s), wash at once with large quantities of water.
Drying
Dry heat-resistant glass pipettes in a hot-air oven at 60 °C and ordinary pipettes in an incubator at 37 °C. Alternatively, leave pipettes to air-dry.
Using the vacuum pump
This is a small instrument made of metal, plastic or glass that is attached to the water tap.
1. Turn the water on hard to drive a strong jet through the pump. This causes air to be sucked into the side arm of the pump and the rubber tubing attached to it.
2. Fit this rubber tubing over the tip of the pipette.
3. Dip the other end of the pipette into the rinsing liquid (water or detergent solution), which is sucked through the pipette and discharged by the pump into the sink (Fig. 3.59).
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| Fig. 3.59 Using a vacuum pump to rinse a pipette |
Microscope slides
New slides
Soaking in detergent solution
Prepare a bowl of water mixed with washing powder or liquid detergent. Use the amounts recommended by the manufacturer. Place the slides in the bowl one by one and leave to soak overnight.
Rinsing in water
Rinse each slide with tap water and then soak in clean water for 15 minutes.
Wiping and drying
Wipe the slides, one at a time, with a soft, non-fluffy cloth. Place them on a sheet of filter paper, one by one. Leave to dry. Examine each slide. Discard any slides that are stained, scratched or yellow or that have dull patches on them, or try to clean them again.
Wrapping up
Divide the slides into piles of 10 or 20 and wrap each pile in a small sheet of paper.
Numbering
In some laboratories the slides are numbered in advance in series of five packets with a diamond pencil. (For example, for packets containing 20 slides each, the slides are numbered 1–20, 21–40, 41–60, 61–80 and 81–100, respectively.)
Dirty slides
Slides covered with immersion oil Take the oily slides one by one and rub them with newspaper to remove as much of the oil as possible.
Slides with coverslips
Using the tip of a needle or forceps, detach the coverslips and drop them into a beaker of water (Fig. 3.60) (for cleaning of coverslips, see overleaf).
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| Fig. 3.60 Removing coverslips from a slide for cleaning |
Soaking in detergent solution
Prepare a bowl of cold or lukewarm water mixed with detergent. Use the amount recommended by the manufacturer to produce a strong detergent solution.
Leave the slides to soak for 24 hours.
Note: Detergents containing enzymes are excellent for removing blood films. When slides have been used for infected specimens (e.g. urine, stools), they should be placed in disinfectant solution before cleaning.
Cleaning
After the slides have soaked for 24 hours, prepare another bowl containing a weak detergent solution (15 ml of household detergent per litre of water).
Remove the slides one by one from the strong detergent solution.
Rub each one with cotton wool dipped in the strong detergent solution, then drop into the bowl of weak detergent solution and leave to soak for 1 or 2 hours.
Rinsing
Preferred method
Remove the slides one by one from the weak detergent solution using forceps. If you must use your fingers, pick the slides up by their edges. Rinse each slide sepa- rately under the tap, then soak for 30 minutes in a bowl of water.
Quick method
Empty the bowl of weak detergent solution and fill with clean water. Change the water three times, shaking the bowl vigorously each time.
Wiping, drying and wrapping up
Follow the instructions given above for new slides.
Coverslips
Used coverslips can be cleaned and reused.
1. Make up the following solution in a large beaker:
— 200 ml of water
— 3 ml of detergent
— 15 ml of bleach or 5 ml of a quaternary ammonium disinfectant.
2. Put the coverslips into the beaker one by one.
3. Leave the coverslips to soak for 2–3 hours, shaking gently from time to time.
4. Rinse out the beaker containing the coverslips with tap water four times, shaking gently.
5. Give a final rinse with demineralized water.
6. Drain the coverslips by tipping them out carefully on to a pad of gauze.
7. Dry in a hot-air oven at 60 °C, if possible.
Keep clean, dry coverslips in a small Petri dish. If possible, use special coverslip forceps for taking them out.
Reusable syringes and needles
As soon as a sample has been collected, remove the plunger from the used syringe and rinse both the barrel and the plunger. Fill the barrel with water, insert the plunger and force the water through the needle. Finally remove the needle and rinse the hub cavity.
Reusable syringe with blocked piston
To loosen the piston, choose one of the following methods:
● Soak for 2 hours in hot water (about 70 °C).
● Stand the syringe on its end, piston down. Pipette 50% acetic acid solution (reagent no. 3) into the nozzle of the syringe with a fine Pas- teur pipette (Fig. 3.61). Leave for 10 minutes.
After loosening the piston, soak the syringe for several hours in a bowl of 1 mmol/l hydrogen peroxide.
.jpg "Cleaning a blocked (reusable) syringe using acetic acid") |
Fig. 3.61 Cleaning a blocked (reusable)
syringe using acetic acid |
Rinsing and soaking needles
As soon as the needle has been used, rinse it while it is still attached to the syringe, then remove it and leave it to soak in hot water.
Blocked needles
To remove the blockage, use a nylon thread dipped in 50% acetic acid solution (reagent no. 3); alternatively, you can use a stylet.
2. Cleaning non-disposable specimen containers
Non-disposable containers, such as jars and bottles, may contain stools, sputum, pus, CSF, blood or urine, all of which may harbour potentially infectious organisms.
Containers for stool specimens
If the lavatory is not connected to a septic tank, fill the jars containing stools with a 5% solution of cresol (see page 83) or a similar disinfectant. Leave for 6 hours.
Empty into the lavatory.
If the lavatory is connected to a septic tank, cresol or other disinfectants should not be added to the stools before disposal. Clean the jars with detergent and water, as described on page 80.
Sputum pots and tubes containing pus or CSF specimens
There are several possible methods.
Using an autoclave
This is the best method.
1. Place the containers in the autoclave and sterilize for 30 minutes at 120 °C.
2. After the containers have cooled, empty the contents into the sink or lavatory.
3. Clean with detergent and water, as described on page 80.
Boiling in detergent
Keep a large pan especially for this purpose.
Boil sputum pots for 30 minutes in water containing washing powder (60 g per litre of water) (Fig. 3.62)
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| Fig. 3.62 Cleaning sputum pots by boiling in detergent |
Using formaldehyde solution or cresol
Pour into each sputum pot either:
— 10 ml of undiluted formaldehyde, 10% solution (reagent no. 28), or
— 5 ml of 5% cresol (see page 83). Leave for 12 hours.
Urine bottles
Empty the bottles into the lavatory. Fill them with either:
— a 10% solution of household bleach (see page 84), or
— a 5% solution of cresol (see page 83). Leave for 4 hours.
Test-tubes containing blood specimens
Test-tubes of fresh blood collected on the same day should be:
— rinsed in cold water
— left to soak in a detergent solution (see page 80).
Test-tubes of “old” blood kept for several days at room temperature may contain large numbers of microorganisms. They should be:
— filled with a 10% solution of household bleach (see page 84)
— left for 12 hours and then
3. Cleaning and maintenance of other laboratory equipment
Centrifuges
Clean the bowl of the centrifuge daily or after any spillage occurs. Use 70% ethanol for metal bowls and 1% bleach (see page 84) for plastic ones. (Do not use bleach for metal bowls as it may cause corrosion.)
Rinse the centrifuge buckets after use and remove any traces of blood, etc.
Check the wiring for fraying and loose connections at regular intervals. If the cen- trifuge is sparking or running irregularly, the carbon brushes may need replacing.
Lubrication of the centrifuge should be carried out by a specialist, according to the manufacturer’s instructions.
Water-baths
If possible fill the water-bath with distilled water or rainwater to prevent deposits forming inside. A crystal of thymol will help to prevent algal growth.
Change the water and clean the inside of the water-bath at least once a month or whenever it looks dirty. Use a thermometer to check the water temperature each time the water is changed as scale on the heating element may cause the thermostat to malfunction.
Incubators
Incubators are used for bacterial culture by laboratories working in microbiology. The incubator must maintain a constant average temperature of 35 °C (range 33–37 °C). The actual temperature must correspond to the thermostat setting when the instrument is used.
In carbon dioxide incubators used for microbial culture, the concentration of carbon dioxide should be maintained at 5–10% and the humidity at 50–100%.
The temperature in the incubators should be recorded daily. Like all laboratory instruments, incubators must be cleaned at regular intervals (at least every fortnight) and also after spillage of any material, whether infectious or non-infectious.
Westergren tubes
Rinse in water, then leave to soak in clean water for 12 hours. Dry completely (in an incubator at 37 °C, if possible). Do not use washing powder, acids or ethanol.
4. Disinfectants
There are many disinfectants that have various different chemical actions on infec- tive agents. Table 3.1 lists the disinfectants that are most commonly used in health laboratories.
Cresols
Cresols may be solid or liquid; they are less water-soluble than phenol, but a 5% aqueous solution can be kept as a stock solution. Cresols emulsify well in soap solutions.
Lysol
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a Chemical disinfection for skin-cutting and invasive instruments should be employed only as the last resort, if neither
sterilization nor high- level disinfection by boiling is
possible, and then only if the appropriate concentration of the chemical is available
and if the instruments have been thoroughly cleaned to remove gross contamination before
soaking in the chemical disinfectant.
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exposure of material to phenol solution must be longer than for cresol. Phenol and cresol solutions cause irritation of the skin and eyes.
Sodium and calcium hypochlorite
Sodium and calcium hypochlorite solutions (household bleaches) are very strong disinfectants. They are used in a number of laboratory, household and industrial applications. Hypochlorites are rapidly inactivated by particles of dust and organic materials and must be freshly prepared from stock solutions every day. Hypochlorites cause irritation of the skin, eyes and lungs.
Strong, undiluted solutions should contain 10% available chlorine.
For preparing working dilutions, the following dilutions are recommended:
● For jars and containers in which used pipettes, slides or other glassware are discarded and for swabbing bench surfaces: 10 ml of concentrated hypochlorite solution in 990 ml of water (0.1% available chlorine). Place the used glassware into the jars of hypochlorite solution and leave for at least 12 hours. Do not overfill the containers. Change the containers daily.
● For decontamination of blood spills and other specimens with a high protein content: 40 ml of concentrated hypochlorite solution in 360 ml of water (1% available chlorine).
Strong hypochlorite solutions are corrosive and can cause burns. Handle solutions of bleach carefully: wear rubber gloves to protect the hands, and eye shields to prevent splashing in the eyes.
Calcium hypochlorite is available in its solid form as powder or granules. It decomposes at a slower rate than sodium hypochlorite. A solution of 1% available chlor- ine is obtained by dissolving 14 g of calcium hypochlorite in 1 litre of water.
Chloramine
Chloramine (tosylchloramide sodium) is a crystalline powder which, like the hypochlorites, releases chlorine as the active disinfectant agent, although at a slower rate. It is also used for water disinfection: chlorinated water has a concentration of 0.05% chloramine. Note that chlorinated water can interfere with laboratory tests. Distilled water must therefore be used.
Calcium hydroxide
Calcium hydroxide solution is prepared from quicklime (calcium oxide) powder or granules dissolved in water (1 part : 3 parts w/v). Calcium hydroxide solution is not suitable for disinfecting stools from patients with tuberculosis.
Quaternary ammonium compounds
Quaternary ammonium compounds (QUATS) are effective against vegetative bacteria and some fungi. They are not effective against spores, viruses and mycobacte- ria; they are not toxic and are harmless to the skin.
Alcohols
Alcohols (e.g. ethanol, isopropanol, n-propanol) are fast-acting, but relatively expensive disinfectants that are usually used for skin disinfection. They kill bacteria and some viruses, but not fungi.
Iodine
Iodine is an excellent, fast-acting disinfectant with a wide range of action. It kills bacteria, many spores, viruses and fungi. At low temperatures iodine is more active than other disinfectants. Some people are hypersensitive to iodine and suffer a rash on areas of skin that have been exposed to iodine solution. Their sensitivity is much less when iodophores (polymer solutions that bind iodine) such as polyvidone iodine are used.
5. Sterilization
Sterilization is defined as the destruction of all microorganisms in or about an object. In the medical laboratory sterilization is achieved either by moist heat (autoclaving, boiling) or by dry heat (hot-air oven, flaming). Materials are steri- lized for three main purposes in the medical laboratory:
— in preparation for taking specimens (needles, syringes, tubes, etc. must be sterile);
— to disinfect contaminated materials;
— to prepare the equipment used for bacteriological cultures (Petri dishes, Pasteur pipettes, tubes, etc.).
Sterilization by steam
Using an autoclave
Clinical samples and other contaminated waste materials are placed in a special autoclave bag or into a metal or plastic bucket for autoclaving. Use the autoclave sterilizing indicators to control the sterilizing cycle.
Principle
Water is heated in a closed container. This produces saturated steam under pres- sure, with a temperature of over 100 °C. Most types of microorganism, including all bacteria (but not all viruses) are killed when apparatus is heated for 20 minutes at 120 °C in this steam under pressure.
Components of an autoclave (Fig. 3.63)
1. Boiler
A large deep cylinder in which the items to be sterilized are placed.
2. Basket
A big wire basket that holds the materials to be sterilized.
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Fig. 3.63 Components of an autoclave
1: boiler; 2: basket; 3: basket support; 4: drainage tap; 5: lid;
6: lid clamps; 7: air outlet valve;
8: safety valve; 9: temperature gauge or pressure gauge. |
3. Basket support
A support in the bottom of the autoclave that holds the basket above the water level.
4. Drainage tap
A tap fitted at the base of the boiler to drain off excess water.
5. Lid
The lid covers and seals the boiler and is fitted with a rubber washer.
6. Lid clamps
These clamps, together with the rubber washer, seal the lid and pre- vent steam from escaping.
7. Air outlet valve
A valve at the top of the boiler or on the lid that is used to let air out when the water is first heated.
8. Safety valve
A valve at the top of the boiler or on the lid that lets steam escape if the pressure becomes too high and so prevents an explosion.
9. Temperature gauge or pressure gauge
All gauges indicate the temperature in degrees Celsius (°C); some also have a second set of figures indicating the pressure.
Heating system
The heating system may be built into the autoclave in the form of:
— electric elements
— gas burners
— a paraffin oil stove.
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Fig. 3.64 Autoclaving syringes and needles
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Installation
Autoclaves should be installed away from the main working area, as they are noisy. If gas or a paraffin oil stove is used for heating, it should be kept away from flammable materials and chemicals.
Preparation of material for sterilization
Reusable syringes
Reusable syringes are placed in large glass test-tubes plugged with non-absorbent cotton wool (the pistons and barrels in separate tubes; Fig. 3.64), or they are wrapped in gauze and placed in metal trays.
Reusable needles
Reusable needles should be placed separately in small test-tubes that are then plugged (see Fig. 3.64). Place a pad of non-absorbent cotton wool at the bottom of each tube to protect the tip of the needle.
Otherwise, arrange the needles in metal trays with their points stuck into a folded gauze pad (Fig. 3.65).
The metal trays are placed uncovered in the autoclave.
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| Fig. 3.65 Alternative method for autoclaving needles |
Glassware
Specimen tubes, Petri dishes, etc. should be wrapped in autoclavable polyethylene bags and tied with string.
Pasteur pipettes (Fig. 3.66)
Pasteur pipettes should be placed in large tubes which are then plugged.
Alternatively they may be placed in autoclavable polyethylene bags.
Sterilization procedure
1. Fill the bottom of the autoclave with water (up to the basket support).
Make sure that the water does not touch the basket. If necessary, drain off excess water by opening the drainage tap.
2. Put the basket containing the material to be sterilized in the autoclave together with sterilization indicator papers. The indicator papers turn black when the correct temperature is reached.
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| Fig. 3.66 Autoclaving Pasteur pipettes |
3. Close the lid, making sure that the rubber washer is in its groove. Screw down the lid clamps evenly and firmly, but not too tightly.
4. Open the air outlet valve.
5. Begin heating the autoclave.
6. Watch the air outlet valve until a jet of steam appears. Wait 3 or 4 minutes until the jet of steam is uniform and continuous. This shows that all the air has been driven out of the autoclave.
7. Close the air outlet valve. Tighten the lid clamps and reduce the heat slightly.
8. Watch the temperature gauge. When the desired temperature is reached (i.e. 120 °C) the heat must be regulated to maintain it. Reduce the heat until the needle on the dial remains at the temperature selected. Start timing at this point.
Sterilization times
● Materials for collecting specimens (reusable syringes and needles, tubes): 20 minutes at 120 °C.
● Containers of infected material (sputum pots, tubes of pus): 30 minutes at 120 °C.
● Bacteriological culture media: follow the instructions of the bacteriologist or the chief laboratory technician.
Turning off the heat
1. Turn off the heat as soon as the required time is up.
2. When the temperature falls below 100 °C, open the air outlet valve to equalize the pressures inside and outside the autoclave.
3. When the hissing sound stops, unscrew the lid clamps. Take off the lid. Leave the autoclave to cool, then carefully remove the basket of sterile equipment. If drops of water have formed, dry the sterile equipment in an incubator at 37 °C, if possible.
Cleaning
Wipe the inside of the autoclave daily or whenever spillages occur.
Precautions
● Never touch the drainage tap, outlet valve or safety valve of the autoclave while heating it under pressure.
● Never heat the autoclave too quickly to bring up the pressure once the outlet valve is closed.
● Never leave the autoclave unattended while the pressure is rising.
● Never open the lid before the pressure has dropped to normal, as you may be scalded with steam.
● During sterilization make sure the lid is secured and no steam escapes as if it does, neither the pressure nor the temperature will be correct.
● Never leave the autoclave to cool for too long, because if it is left for several hours without the outflow valve being opened, a vacuum forms.
Using a pressure cooker
Pressure cookers are large saucepans designed to cook food very quickly, using steam under pressure. They are used in some small laboratories to sterilize equip- ment used for specimen collection.
Pressure cooker with revolving valve
1. Fill the bottom of the pressure cooker with water. Place the material or object to be sterilized in the basket (which is held above the water level by a support). The wrapped articles should be placed upright (never lay them flat; Fig. 3.67).
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Fig. 3.67 Sterilizing equipment using a pressure cooker
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2. Fit on the lid. Screw it down with its knob. Place the revolving valve (V1) on its shaft in the lid (Fig. 3.68).
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Fig. 3.68 Components of a
pressure cooker
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3. Start heating the pressure cooker on the stove. The valve soon begins to turn, letting a jet of steam escape.
4. Wait until the jet of steam is continuous, then lower the heat so that the valve keeps turning slowly. Leave the pressure cooker on moderate heat for 20 minutes.
5. Turn off the heat. Leave the pressure cooker to cool (or cool it in cold water).
6. Pull off the revolving valve so that air can enter. Remove the lid. Take out the sterilized material or object and leave the pressure cooker to dry.
Warning: Never touch the safety valve (V2 in Fig. 3.68), which is fixed to the lid.
Sterilization by boiling
This method should be used only where there is no alternative. Use a special boil- ing pan or, if not available, a saucepan. Fill the pan with water (preferably demineralized) and heat over the stove. Glassware (reusable syringes) should be put in while the water is still cold. Metal articles (reusable needles, forceps) should be put in when the water is boiling. Leave the articles to boil for 30 minutes.
Sterilization by dry heat
Using a hot-air oven
This method should be used only for glass or metal articles (reusable syringes and needles, pipettes, etc.) when an autoclave is not available. It must not be used for culture media used in microbiology, which should be autoclaved (see page 86).
1. Prepare the object to be sterilized in the same way as for the autoclave method.
Cotton-wool plugs should not be too thick, otherwise the hot air cannot penetrate. Raise the lids of the metal boxes slightly and arrange them so that they face the back of the oven.
2. Set the thermostat to 175 °C and switch on the oven. If there is a fan, check that it is working.
3. Watch the thermometer. When the temperature reaches 175 °C, continue heat- ing at this temperature for a further 60 minutes. If the object to be sterilized is heavy or bulky or if it includes powders, oils or petroleum jelly, heat at 175 °C for 2 hours.
4. Switch off the heat. Wait until the temperature falls to 40 °C. Open the oven door. Close the lids of the metal boxes. Remove the sterile object.
By flaming
This method should be used only for metal articles such as for- ceps and scalpels. It is not suitable for general use.
1. Place the articles in a metal tray.
2. Add about 10 drops of ethanol and ignite.
3. During flaming tilt the tray first one way, then the other (Fig. 3.69).
To sterilize bacteriological loops, heat them in the flame of a gas burner or spirit lamp until they are red hot.
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| Fig. 3.69 Sterilization by flaming |
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