Molecular detection methods have the ability to produce a large volume of nucleic acid through the amplification of trace quantities found in samples. While this is beneficial for enabling sensitive detection, it also introduces the possibility of contamination through the spreading of amplification aerosols in the laboratory environment. When conducting experiments, measures can be undertaken to avoid the contamination of reagents, laboratory equipment and bench space, as such contamination may generate false-positive (or false-negative) results.
To help reduce the likelihood of contamination, Good Laboratory Practice should be exercised at all times. Specifically, precautions should be taken regarding the following points:
1. Handling reagents
2. Organization of workspace and equipment
3. Use and cleaning advice for the designated molecular space
4. General molecular biology advice
5. Internal controls
6. Bibliography
1. Handling reagents
Briefly centrifuge reagent tubes before opening to avoid the generation of aerosols. Aliquot reagents to avoid multiple freeze-thaws and the contamination of master stocks. Clearly label and date all reagent and reaction tubes and maintain logs of reagent lot and batch numbers used in all experiments. Pipette all reagents and samples using filter tips. Prior to purchase, it is advisable to confirm with the manufacturer that the filter tips fit the brand of pipette to be used.
2. Organization of workspace and equipment
Workspace should be organized to ensure that the flow of work occurs in one direction, from clean areas (pre-PCR) to dirty areas (post-PCR). The following general precautions will help to reduce the chance of contamination. Have separate designated rooms, or at minimum physically separate areas, for: mastermix preparation, nucleic acid extraction and DNA template addition, amplification and handling of amplified product, and product analysis, e.g. gel electrophoresis.
In some settings, having 4 separate rooms is difficult. A possible but less desirable option is to do the mastermix preparation in a containment area, e.g. a laminar flow cabinet. In the case of nested PCR amplification, the preparation of the mastermix for the second round reaction should be prepared in the ‘clean’ area for mastermix preparation, but the inoculation with the primary PCR product should be done in the amplification room, and if possible in a dedicated containment area (e.g. a laminar flow cabinet).
Each room/area needs a separate set of clearly labelled pipettes, filter tips, tube racks, vortexes, centrifuges (if relevant), pens, generic lab reagents, lab coats and boxes of gloves that will remain at their respective workstations. Hands must be washed and gloves and lab coats changed when moving between the designated areas. Reagents and equipment should not be moved from a dirty area to a clean area. Should an extreme case arise where a reagent or piece of equipment needs to be moved backwards, it must first be decontaminated with 10% sodium hypochlorite, followed by a wipe down with sterile water.
Note
The 10% sodium hypochlorite solution must be made up fresh daily. When used for decontamination, a minimum contact time of 10 minutes should be adhered to.
Alternatively, commercially available products that are validated as DNA-destroying surface decontaminants can be used if local safety recommendations do not allow the use of sodium hypochlorite or if sodium hypochlorite is not suitable for decontaminating the metallic parts of equipment.
Ideally, staff should abide by the unidirectional work flow ethos and not go from dirty areas (post-PCR) back to clean areas (pre-PCR) on the same day. However, there may be occasions when this is unavoidable. When such occasion arises, personnel must take care to thoroughly wash hands, change gloves, use the designated lab coat and not introduce any equipment they will want to take out of the room again, such as lab books. Such control measures should be emphasized in staff training on molecular methods.
After use, bench spaces should be cleaned with 10% sodium hypochlorite (followed by sterile water to remove residual bleach), 70% ethanol, or a validated commercially available DNA-destroying decontaminant. Ideally, ultra-violet (UV) lamps should be fitted to enable decontamination by irradiation. However, the use of UV lamps should be restricted to closed working areas, e.g. safety cabinets, in order to limit the laboratory staff’s UV exposure. Please abide by manufacturer instructions for UV lamp care, ventilation and cleaning in order to ensure that lamps remain effective.
If using 70% ethanol instead of sodium hypochlorite, irradiation with UV light will be needed to complete the decontamination.
Do not clean the vortex and centrifuge with sodium hypochlorite; instead, wipe down with 70% ethanol and expose to UV light, or use a commercial DNA-destroying decontaminant. For spills, check with the manufacturer for further cleaning advice. If manufacturer instructions permit it, pipettes should be routinely sterilized by autoclave. If pipettes cannot be autoclaved, it should suffice to clean them with 10% sodium hypochlorite (followed by a thorough wipe down with sterile water) or with a commercial DNA-destroying decontaminant followed by UV exposure.
Cleaning with high-percentage sodium hypochlorite may eventually damage pipette plastics and metals if done on a regular basis; check recommendations from the manufacturer first. All equipment needs to be calibrated regularly according to the manufacturer-recommended schedule. A designated person should be in charge of ensuring that the calibration schedule is adhered to, detailed logs are maintained, and service labels are clearly displayed on equipment.
3. Use and cleaning advice for the designated molecular space
Pre-PCR: Reagent aliquoting / mastermix preparation: This should be the cleanest of all spaces used for the preparation of molecular experiments and should ideally be a designated laminar flow cabinet equipped with a UV light. Samples, extracted nucleic acid and amplified PCR products must not be handled in this area. Amplification reagents should be kept in a freezer (or refrigerator, as per manufacturer recommendations) in the same designated space, ideally next to the laminar flow cabinet or pre-PCR area. Gloves should be changed each time upon entering the pre-PCR area or laminar flow cabinet.
The pre-PCR area or laminar flow cabinet should be cleaned before and after use as follows: Wipe down all items in the cabinet, e.g. pipettes, tip boxes, vortex, centrifuge, tube racks, pens, etc. with 70% ethanol or a commercial DNA-destroying decontaminant, and allow to dry. In the case of a closed working area, e.g. a laminar flow cabinet, expose the hood to UV light for 30 minutes.
Note
Do not expose reagents to UV light; only move them into the cabinet once it is clean. If performing reverse transcription PCR, it may also be helpful to wipe down surfaces and equipment with a solution that breaks down RNases on contact. This may help to avoid false-negative results from enzyme degradation of RNA. After decontamination and before preparing the mastermix, gloves should be changed once more, and then the cabinet is ready to use.
Pre-PCR: Nucleic acid extraction/template addition:
Nucleic acid must be extracted and handled in a second designated area, using a separate set of pipettes, filter tips, tube racks, fresh gloves, lab coats and other equipment.This area is also for the addition of template, controls and trendlines to the mastermix tubes or plates. To avoid contamination of the extracted nucleic acid samples that are being analysed, it is recommended to change gloves prior to handling positive controls or standards and to use a separate set of pipettes. PCR reagents and amplified products must not be pipetted in this area. Samples should be stored in designated fridges or freezers in the same area. The sample workspace should be cleaned in the same way as the mastermix space.
Post-PCR: Amplification and handling of the amplified product
This designated space is for post-amplification processes and should be physically separate from the pre-PCR areas. It usually contains thermocyclers and real-time platforms, and ideally should have a laminar flow cabinet for adding the round 1 PCR product to the round 2 reaction, if nested PCR is being performed. PCR reagents and extracted nucleic acid must not be handled in this area since the risk of contamination is high. This area should have a separate set of gloves, lab coats, plate and tube racks, pipettes, filter tips, bins and other equipment. Tubes must be centrifuged before opening. The sample workspace should be cleaned in the same way as the mastermix space.
Post-PCR: Product analysis
This room is for product detection equipment, e.g. gel electrophoresis tanks, power packs, UV transilluminator and the gel documentation system. This area should have separate sets of gloves, lab coats, plate and tube racks, pipettes, filter tips, bins and other equipment. No other reagents can be brought into this area, excluding loading dye, molecular marker and agarose gel, and buffer components. The sample workspace should be cleaned in the same way as the mastermix space.
Important note
Ideally, the pre-PCR rooms should not be entered on the same day if work has already been performed in the post-PCR rooms. If this is completely unavoidable, ensure that hands are first washed thoroughly and that specific lab coats are worn in the rooms. Lab books and paperwork must not be taken into the pre-PCR rooms if they have been used in the post-PCR rooms; if necessary, take duplicate print-outs of protocols/sample IDs, etc.
4. General molecular biology advice
Use powder-free gloves to avoid assay inhibition. Correct pipetting technique is paramount to reducing contamination. Incorrect pipetting can result in splashing when dispensing liquids and the creation of aerosols. Good practice for correct pipetting can be found at the following links: Gilson guide to pipetting, Anachem pipetting technique videos, Centrifuge tubes before opening, and open them carefully to avoid splashing. Close tubes immediately after use to avoid the introduction of contaminants.
When performing multiple reactions, prepare one mastermix containing common reagents (e.g. water, dNTPs, buffer, primers and enzyme) to minimize the number of reagent transfers and reduce the threat of contamination. It is recommended to set up the mastermix on ice or a cold block. Use of a Hot Start enzyme may help to reduce the production of non-specific products. Protect reagents containing fluorescent probes from light in order to avoid degradation.
5. Internal controls
Include well-characterized, confirmed positive and negative controls, along with a no-template control in all reactions, and a multi-point titrated trendline for quantitative reactions. The positive control should not be so strong that it poses a contamination risk. Include positive and negative extraction controls when performing nucleic acid extraction.
It is recommended that clear instructions be posted in each of the areas so that users are aware of rules of conduct. Diagnostic labs detecting very low levels of DNA or RNA in clinical samples may want to adopt the additional security measure of having separate air handling systems with slightly positive air pressure in the pre-PCR rooms and slightly negative air pressure in the post-PCR rooms.
Lastly, developing a quality assurance (QA) plan is helpful. Such a plan should include lists of reagent master stocks and working stocks, rules for storing kits and reagents, reporting of control results, staff training programmes, troubleshooting algorithms, and remedial actions when needed.
6. Bibliography
Aslan A, Kinzelman J, Dreelin E, Anan’eva T, Lavander J. Chapter 3: Setting up a qPCR laboratory. A guidance document for testing recreational waters using USEPA qPCR method 1611. Lansing- Michigan State University.
Public Health England, NHS. UK standards for microbiology investigations: Good Laboratory Practice when performing molecular amplification assays). Quality Guidance. 2013;4(4):1–15.
Mifflin T. Setting up a PCR laboratory. Cold Spring Harb Protoc. 2007;7.
Schroeder S 2013. Routine maintenance of centrifuges: cleaning, maintenance and disinfection of centrifuges, rotors and adapters (White paper No. 14). Hamburg: Eppendorf; 2013.
Viana RV, Wallis CL. Good Clinical Laboratory Practice (GCLP) for molecular based tests used in diagnostic laboratories, In: Akyar I, editor. Wide spectra of quality control. Rijeka, Croatia: Intech; 2011: 29–52.
Post time: Jul-16-2020