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Screening for the hepatitis C virus in some dental clinics in Alexandria, Egypt

Hashish, Mona H.a; Selim, Heba S.a; Elshazly, Soraya A.a; Diab, Hanan H.b; Elsayed, Naguiba M.b

The Journal Of The Egyptian Public Health Association: December 2012 - Volume 87 - Issue 5 and 6 - p 109–115
doi: 10.1097/01.EPX.0000421670.02166.ec
Original articles
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Background/objectives Individuals can be exposed to the hepatitis C virus (HCV) infection through inadequately or improperly sterilized medical or dental equipment. The aim of this study was to detect HCV RNA in the dental setting in Alexandria, Egypt.

Materials and methods The study included 100 samples collected from five dental clinics (A–E) in Alexandria. The samples were collected from critical, semicritical, and noncritical instruments during different periods of the day (morning, mid-day, end of the day). Samples were subjected to a reverse transcriptase-PCR for the detection of HCV RNA.

Results HCV RNA was detected in 18% (18 out of 100) of the instrument samples tested. Two positive HCV RNA samples were collected from semicritical instruments in clinic B, whereas 16 positive HCV RNA samples were collected from clinic D (eight samples from critical, six samples from semicritical, and two samples from noncritical instruments). There was a statistically significant difference between clinics B and D in terms of the samples collected in the morning and those collected at the end of the day.

Conclusion and recommendations HCV RNA as detected by PCR was found in a considerable percent of instruments’ samples (18%). Most of the positive HCV RNA samples (16 out of 18 samples) obtained from instruments were among those collected from clinic D. This clinic used only glutaraldehyde as a method of sterilization. Therefore, proper infection control measures, including sterilization and disinfection should be strictly adopted.

aDepartment of Microbiology, High Institute of Public Health

bDepartment of Oral Medicine, Faculty of Dentistry, Alexandria University, Alexandria, Egypt

Correspondence to Mona H. Hashish, Department of Microbiology, High Institute of Public Health, Alexandria University, 165 Elhorreya Str. Elhadara, Alexandria, Egypt Tel: +2 01223525401; e-mail: monash64@yahoo.com

Received September 13, 2012

Accepted September 18, 2012

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Introduction

Hepatitis C virus (HCV) infection represents a major public health problem in the worldwide currently. The WHO has estimated a worldwide prevalence of about 3%, with the virus affecting more than 170 million individuals 1. Egypt has the highest countrywide prevalence of HCV in the world, with an estimated 10–15% antibody prevalence rate 2.

Transmission occurs primarily by percutaneous exposure to contaminated blood or saliva. In clinical institutions, medical and many dental instruments may come into contact with patients’ body fluids such as saliva, blood, or mucous membranes during treatment or examination. Hence, proper sterilization of invasive instruments and equipments is very important in the practice of dentistry, where various blood-borne or saliva-borne pathogens could be easily transmitted to patients and dental staff through contaminated or inadequately sterilized instruments 3. The American Dental Association (ADA) recommends that surgical and other instruments that normally penetrate soft tissue or bone (e.g. extraction forceps, scalpel blades, bone chisels, periodontal scalers, and surgical burs) be classified as critical devices that should be sterilized after each use or discarded. Instruments not intended to penetrate oral soft tissues or bone (e.g. amalgam condensers and air/water syringes) but that could contact oral tissues are classified as semicritical, but sterilization after each use is recommended if the instruments are heat-tolerant 4.

Autoclaving is an effective method for virus destruction, but may not be available in all rural or suburban areas 5. Dry heat sterilization is used for materials that might be damaged by moist heat (e.g. burs); however, it is a prolonged process and the high temperatures required are not suitable for certain patient care items and devices. Heat-sensitive critical and semicritical instruments can be sterilized by immersion in liquid chemical germicides (e.g. glutaraldehyde, peracetic acid, and hydrogen peroxide). Manufacturer instructions must be strictly followed when using these chemical sterilants in terms of dilution, immersion time, and temperature. In addition, poststerilization procedures are essential where items need to be rinsed with sterile distilled water, handled using sterile gloves, and dried with sterile towels 6. According to the Centers for Disease Control and Prevention (CDC), noncritical contact surfaces are surfaces that might be touched frequently with gloved hands during patient care or that might become contaminated with blood or other potentially infectious material and subsequently contact instruments, hands, gloves, or devices (e.g. light handles, switches, dental radiograph equipment, chair-side computers). Barrier protective coverings (e.g. clear plastic wraps) can be used for these surfaces, particularly those that are difficult to clean (e.g. light handles, chair switches) 7.

Dental treatment has been reported to be the only risk factor by some HCV-positive patients 8. Thus, the aim of this study was to screen for HCV in relation to infection control measures in five dental clinics in Alexandria (Egypt).

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Materials and methods

This cross-section study was carried out in five (two private and three educational) dental clinics in Alexandria (A, B, C, D, and E) during the year 2010. The clinics were inspected in terms of the methods of sterilization used, the presence of an instrument-processing area, washing of instruments, wrapping for the dental unit between patients, and the usage of gloves. Based on a pilot study, where HCV infection was detected in 8% of non sterile instruments, and assuming 0% detection rate among sterile instruments; at a power of 80% and a level of 0.05, the minimum required sample size was calculated to be 94 using Minitab Version 13. Samples were collected on three occasions during the day (morning, mid-day, end of the day) from sterile instruments, used instruments, and unused instruments. One hundred dental setting samples (20 from each clinic) were collected from the following categories:

  • Surgical instruments (e.g. forceps, bone file, scalpel, surgical bur, needle holder).
  • Diagnostic instruments (e.g. mirrors, probes, tweezers).
  • Operative instruments (e.g. endodontic files, burs both diamond and carbide, burnishers).
  • Orthodontic instruments.
  • High-speed and low-speed handpieces.
  • Parts of the dental chair (e.g. bracket table handle, switches of the bracket table, suction tip, and spittoon).

The above-mentioned patient-care items were categorized as critical, semicritical, or noncritical, depending on the potential risk for infection associated with their intended use. For sterilization of both critical and noncritical instruments of the five clinics, two clinics (A and E) had an autoclave, two had both an autoclave and a hot-air oven (B and C), and one clinic (D) used glutaraldehyde for sterilization because of the breakdown of the autoclave they had. All the clinics used 50% sodium hypochlorite (bleach) for clinic surfaces.

For each sample, 1 ml sterile PBS was distributed in a sterile 1.5 ml microcentrifuge tube, dipped previously in 0.1% diethylpyrocarbonate for 2 h and sterilized by autoclaving. After immersion in the PBS, the sterile swab was used to wipe instrument surfaces and other parts of the dental clinic (e.g. dental chair, spittoon, and bracket table). The swab was placed back in the PBS and transferred on the same day to the laboratory to be subjected to RNA extraction and reverse transcription steps, followed by storage at −20°C until used for HCV RNA detection by nested-PCR 3. The Z test of proportion was used to compare proportion of positive samples between two clinics, at confidence limit of 95%.

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Detection of hepatitis C virus RNA by nested-polymerase chain reaction

The protocol for HCV RNA detection by nested reverse transcriptase (RT)-PCR amplification proceeds in a stepwise manner from RNA isolation to RT-PCR amplification using external and internal pairs of primers to detection on an agarose gel. RNA was extracted from swab samples by the spin column extraction method using the SV total RNA isolation system (Promega, Madison, Wisconsin, USA) according to the manufacturer’s instructions. HCV RNA was detected by amplification of the 256-bp 5′-untranslated region fragment with gene-specific primers 9.

For the first round of PCR:

External sense primer: 5′-ACT GTC TTC ACG CAG AAA GCG TCT AGC CAT-3′

External antisense primer: 5′-CGA GAC CTC CCG GGG CAC TCG CAA GCA CCC-3′

For the second round of PCR:

Internal sense primer: 5′-ACG CAG AAA GCG TCT AGC CAT GGC GTT AGT-3′

Internal antisense primer: 5′-TCC CGG GGC ACT CGC AAG CAC CCT ATC AGG-3′

Synthesis of cDNA and the first round of PCR were carried out on 10 μl of extracted RNA using the Qiagen OneStep RT-PCR Kit (Qiagen, Hilden, Germany). A volume of 1 μl (1 pmol/μl) of the external antisense primer was used for the synthesis of cDNA using Omniscript and Sensiscript Reverse Transcriptases contained in the kit. Reverse transcription was performed at 50°C for 20 min. 1 μl (1 pmol/μl) of the external sense primer was added to each tube before proceeding to the initial PCR activation step: 95°C for 15 min to activate HotStarTaq DNA polymerase and to simultaneously inactivate the reverse transcriptases. This was followed by 35 cycles, each cycle consisting of 30 s at 95°C, 1 min at 50°C, and 40 s at 72°C, with a final extension period of 7 min at 72°C. For the second round of amplification, 10 μl of the first round PCR product was transferred to a fresh tube containing 40 μl ml of the reaction mixture [12.5 μl Go Taq Green Master Mix (Promega), 0.5 μl RNase-free water, and 1 μl of each of internal sense and antisense primers] and subjected to 35 cycles of amplification, according to the same protocol as the first round of PCR after an initial PCR activation step for 3 min at 94°C. Nuclease-free water as a negative control and a positive HCV sample determined quantitatively by real-time PCR [COBAS AmpliPrep/COBAS TaqMan Real-time PCR (Roche Diagnostics Gmbh, Mannheim, Germany)] were included in each run of PCR. Amplified products were examined after electrophoresis on a 2% agarose gel (Bio Basic Inc., Ontario, Canada) containing 0.5 μg ethidium bromide/ml.

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Results

The infection control measures adopted in the five dental clinics are shown in Table 1. For sterilization, an autoclave was used in clinics A and E, whereas an autoclave and a hot-air oven were used in clinics B and C. In clinic D, glutaraldehyde was the only method of sterilization used. For disinfection, 50% sodium hypochlorite was used in all clinics. In all five clinics, the staff performed washing for the instruments after usage with a special brush, but did not use the utility gloves during manipulation (used normal latex gloves). None of the clinics performed wrapping of the dental chair. Instrument-processing areas were present only in clinics A, C, and E.

Table 1

Table 1

Out of 100 samples tested for the presence of HCV RNA by PCR, 18 (18%) were found to be positive. Two positive HCV RNA results were obtained from instruments’ samples taken from clinic B, 16 positive PCR results were obtained among the instruments’ samples taken from clinic D, and no HCV RNA was detected in any of the tested samples from clinics A, C, and E.

Positive HCV RNA PCR results for the critical and semicritical instruments according to their method of sterilization in the five dental settings are shown in Table 2. Clinics A, B, and E had no positive results for the critical instruments; they were using an autoclave with or without a hot-air oven. For clinic B, the two positive results were found for semicritical instruments despite using the autoclave. In clinic D, 14 positive results were found for critical and semicritical instruments (eight were critical and six were semicritical), where glutaraldehyde was the only means of sterilization.

Table 2

Table 2

In terms of the results of samples from noncritical instruments, not subjected to sterilization (cleaning only), clinics A, B, C, and E were negative for HCV RNA PCR results. The staff used 50% sodium hypochlorite and the clinics were cleaned regularly. Special care was taken for the cleaning of continuously blood contaminated areas (e.g. spittoon, suction tips, and bracket tables). Two positive results were found in clinic D, where the staff admitted cleaning only in the morning and at the end of the day. No cleaning was performed between each patient.

The types of items that were positive for HCV RNA in both clinics B and D are shown in Table 3. In clinic B, which used an autoclave and a hot-air oven for sterilization, a diagnosis mirror and a dental anesthesia syringe were found to be positive. In clinic D, where only glutaraldehyde was used for sterilization, 16 positive samples were obtained from three burs (diamond and carbide, surgical bur), three files (two K files and one H file), one bone file, one scalar tip, one orthodontic plier, one high-speed handpiece, one probe, two dental anesthesia syringes, a rubber dam, a premolar clamp, a suction tip, and a bracket table. The difference between positive samples for clinics B and D was statistically significant (z=6.26, P<0.001).

Table 3

Table 3

Considering the time of sampling, both the positive samples obtained from clinic B were collected at mid-day (one was used and the other was not). Among the 16 positive samples obtained from clinic D, seven were nonused sterile instruments collected during the morning, four were used instruments collected at mid-day, one was a nonused instrument collected at mid-day, and four were used instruments collected at the end of the day. There was a statistically significant difference between clinics B and D in the samples collected in the morning and those collected at the end of the day (Table 4).

Table 4

Table 4

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Discussion

Thousands of individuals infected with the life-threatening HCV may have caught it during routine dental treatment. Hemorrhaging is usually frequent during dental treatment such as oral surgery and scaling. Therefore, contamination of instruments by HCV may occur. Health campaigners have warned that current practices in dental surgery, including the way tools are sterilized, may not be rigorous enough to remove the risk of transmission of the highly infectious virus between patients 10,11.

Infection control procedures, including proper sterilization of invasive instruments are essential to modern dentistry and have an impact on all clinical practices. Accordingly, the present work aimed to investigate the correct and accurate cleaning and decontamination of the dental unit (surfaces touched by both the dentist and the patient) and the adequate sterilization of dental instruments through screening for HCV in sterile and used instruments.

In this study, autoclaving was the main method of sterilization, used in four out of the five studied clinics. Sterilization was performed at a temperature of 120°C for 45 min. Hot-air ovens were used in two of the five clinics at 180°C for 80 min as a method of sterilization besides an autoclave. Clinic D was the only one that used glutaraldehyde as a chemical sterilant. For maximum efficacy of glutaraldehyde as a sterilant, instruments should be immersed in the correct concentration for 9 h. This was hardly followed by the staff of clinic D. Some instruments were only immersed for half an hour or so, and considered to be sterile, and then reused for another patient.

The general causes of sterilization failure include improper packaging, overloading the sterilizer, incorrect operation of the unit, timing errors, mistaking the control for the test strip, and not placing the test strip in the sterilizer unit 12. In addition, some surgical instruments that are used on dental patients have complex narrow channels into which disinfectants and steam penetrate with difficulty 13. Moreover, multiple-use instruments accumulate biofilms of organic residue on their surfaces, and these can protect microorganisms from chemical and physical inactivation. Consequently, sterilization procedures may fail and infection may spread from patient to patient 14,15.

The Ohio Dental State Board has recommended that impervious backed paper, aluminum foil, or plastic wrap must be used to wrap surfaces or items that may be contaminated by blood or saliva and that are difficult or impossible to disinfect. Surface wraps cannot be disinfected and, therefore, if used, must be changed between patients. Surface covers also reduce the handling of chemical disinfectants and require less time to use. When using surface wraps, precleaning and disinfection at the beginning and at end of the day is adequate 16. All aspirators, drains, and spittoons should be cleaned after every session with a surfactant/detergent (to breakdown the biofilms) and a nonfoaming disinfectant 17.

None of the clinics included in the present study performed wrapping of the dental unit. Also, cleaning between the patients was not performed. This may be attributed to the high patient rates, financial abilities of the clinics, and lack of the experience of the dental assistants in infection control procedures.

For surface cleaning, the guidelines of infection control in 2003 stated that bleach should be mixed with water at a dilution of 1–10 or 21 : 100 of a 5.25% solution. A 1 : 10 solution should be used when blood and debris are present. A fresh solution should be prepared every day and heavy utility gloves should be worn 7. In this study, the five dental clinics were unclear about the accurate concentrations of bleach to be used for surface cleaning (noncritical instruments). Mainly a 50% concentration of sodium hypochlorite (bleach) was used. However, no fresh solution was prepared on a daily basis.

Correct decontamination is an essential step in preparing dental instruments for sterilization. Cleaning reduces the bioburden (microorganisms, blood, saliva, oral hard and soft tissues, and dental materials) present. There are two ways to clean dental instruments: mechanically (ultrasonic cleaning, instrument washers, and instrument washer-disinfectors) and manually. If performed well, the level of remaining contamination is low. Sterilization should readily neutralize these moderate amounts of bioburden 18. According to the Ohio Dental State Board, manual cleaning of dental instruments is the least efficient cleaning method. However, if this method is used, the instruments should be fully immersed into a sink prefilled with warm water and detergent and a long-handled kitchen-type brush should be used to remove debris. Instruments should be washed under water with the sharp end of the instrument held away from the body; extra care must be taken when cleaning instruments that are sharp at both ends. Thick waterproof household gloves must be worn to protect against accidental injury and protective eyewear to shield against splashing. The brush used to remove debris from the instruments should be cleaned and autoclaved at regular intervals – at the end of each clinical session, for example. Cleaned brushes should be stored dry 16.

Only manual cleaning was performed in the clinics under study. During washing, the dental assistant used the usual latex glove in all five clinics. The staff were not aware of heavy utility gloves or protective eyewear. The same sink used for instruments was also used by the staff for hand washing in clinics B, D, and E. None of the staff of the five clinics knew how to take care of the brush after usage. Usually, the brush was stored wet.

In this study, 100 samples were collected from different types of instruments (including burs, files, dental anesthesia syringes, dental surgical instruments, orthodontic wires, pliers, high-speed and low-speed handpieces) and some surfaces in the dental clinics. Out of these samples, 18 (18%) were positive for HCV RNA by the PCR technique. Sixteen samples positive for HCV RNA were collected from clinic D. Interestingly, seven (43.75%) HCV RNA-positive samples were collected in the morning from supposedly sterile unused instruments, whereas four (25%) were collected at middle and at the end of the day each. These were used instruments, suggesting contamination by HCV of dental surgeries after the treatment of anti-HCV patients, and that if sterilization and disinfection are inadequate, there is a possible risk of transmission to susceptible individuals. The remaining HCV RNA-positive sample (6.25%) was collected from sterile unused instruments at mid-day, indicating contamination of sterile instruments. The working staff confirmed that during the day, because of the high rate of patients, instruments were only manually cleaned and placed in 2% glutaraldehyde. In clinic B, the two HCV RNA-positive samples were collected at mid-day. One was taken from the diagnosis plate (unused) because of the reuse of the mirrors between patients, whereas the second sample was from a used anesthesia syringe.

In Italy, Piazza et al. 11 have investigated the possible instrument and environmental contamination by HCV RNA in dental surgeries after treatment of anti-HCV-positive and HCV RNA-positive patients. Twenty (6.1%) out of 328 collected samples were positive for HCV RNA. The positive samples were from work benches (two), air turbine handpieces (one), holders (four), suction units (one), forceps (four), dental mirrors (two), and burs (six). The authors found extensive contamination by HCV of dental surgeries after the treatment of anti-HCV patients, necessitating the application of strict infection control measures.

Vickery et al. 19 have examined several sterilization procedures: cleaning in an ultrasonic bath, soaking in 2% gluteraldehyde, autoclaving at 134°C for 3 min, and autoclaving at 121°C for 15 min. They showed that for an anesthetic syringe with a narrow needle lumen, neither ultrasonication nor 2% glutaraldehyde led to effective sterilization. Autoclaving is far more effective, but it is still not reliable for anesthetic syringes, even though the same autoclaving conditions completely inactivate the virus, which is more fully accessible to penetration by steam.

Smith et al.20 have questioned the effectiveness of cleaning schemes on reprocessed endodontic files. Two cleaning regimens dominated: manual cleaning with a brush, followed by autoclaving, and manual cleaning, followed by ultrasonic cleaning and autoclaving. The latter technique is the one recommended by most file manufacturers. Most cleaning regimens, especially those relying primarily on manual scrubbing, proved inadequate. No attempt was made to determine the number of times a given file was used. Therefore, cumulative buildup of contamination was possible. Many of the files were found to be contaminated even without magnification; yet, offices accepted these files for reuse. The authors recommended that the single use of files would eliminate this problem. However, proper cleaning of reusable instruments, followed by correct sterilization are essential infection control measures in dentistry. They also stressed on the fact that endodontic files, rasps, drills, and burs are difficult to clean universally. Thus, such items are best considered as single-use and disposable.

All five clinics investigated in the present work reported that files were thrown only if they were bent or broken. Also, sometimes, the staff did not perform sterilization of files and reamers for fear of breakage or corrosion, or because of the high patient rate. This was also true for burs. This may be the reason for the positive results with such items.

There continues to be some debate about the effective decontamination of handpieces. In theory, a vacuum-phase autoclave will remove the air from the lumen of a dental handpiece, allowing steam to penetrate. The presence of lubricating oil, however, may compromise the sterilization process. Handpieces must be cleaned and autoclaved after each patient 17. The CDC has stated that all permanent handpieces can be contaminated internally with patient material and should be heat sterilized after each patient. Handpieces that cannot be heat sterilized should not be used 5.

In clinic D, one high-speed handpiece yielded a positive result. The staff stated that it is generally known that autoclaving ruins the handpiece. Also, because of the high cost, a clinic cannot afford several handpieces. Thus, handpieces, even with the burs still attached to them, were used from patient to patient.

Thorough cleansing and sterilization of multiuse burs is highly recommended. Cleaning includes presoaking, hand scrubbing, or ultrasonic cleaning and then drying before packaging for the sterilization process. Furthermore, many manufacturers recommend that diamond burs be run against a sharpening stone before ultrasonic cleaning or that the tips be scrubbed with a wire brush to remove any organic debris before sterilization 21. Because these procedures are time consuming and the ADA recommends the use of disposable items whenever possible, the clinical use of disposable burs may be advantageous 22.

In the current work, the reuse of burs between patients was always observed and accompanied by the lack of bur sterilization because of fear of their corrosion. This was mainly because of cost factors. Burs and reamers are expensive and are essential in all dental procedures. This may explain the positive results for the burs found in clinic D.

Generally, many problems may be encountered during the decontamination and sterilization process of dental instruments. Zhou et al. 3 have reported many problems in the decontamination process, including the fact that the instructions for sterilization and disinfection are not always strictly followed. This has been attributed to many factors such as the huge number of patients seeking medical attention daily, low doctor to patient ratio, limited medical equipments, and lack of technical knowledge of the decontamination processes. These factors have resulted in the inadequate sterilization of dental instruments and hence augmented the risk of transmission of microorganisms.

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Conclusion and recommendations

HCV RNA as detected by PCR was found in a huge proportion of instruments’ samples (18%). Most of the positive HVC RNA samples (16 out of 18 samples) obtained from instruments were among those collected from clinic D. This clinic only used glutaraldehyde as a method of sterilization. These results mandate the adoption of strict infection control measures, including sterilization and disinfection. In addition, continuous health education for the dental health care workers in terms of these measures needs to be implemented. Furthermore, dental clinics should have an instrument-processing area with a special basin for instrument wash; in addition, wrapping of the dental unit should be carried out in dental clinics.

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Acknowledgements

Conflicts of interest

There are no conflicts of interest.

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References

1. Worldwide statistics for HCV. Weekly Epidemiological Record No. 49, 10 December 1999, HCV Advocate, Updated Hepatitis C Support Project 2005. Available at: http://www.hcvadvocate.org/hepatitis/hepC/whostats_99.htm [Accessed 12 August 2012]
2. El-Zanaty F, Way A Knowledge and prevalence of hepatitis C. Egypt Demographic and Health Survey 2008. 2009 Cairo Egyptian Ministry of Health, El-Zanaty and Associates, and Macro International:252
3. Zhou LF, Zhu HH, Lin J, Hu MJ, Chen F, Chen Z. Surveillance of viral contamination of invasive medical instruments in dentistry. J Zhejiang Univ Sci B. 2006;7:745–748
4. . Infection control recommendations for the dental office and the dental laboratory. J Am Dent Assoc. 1996;127:672–680
5. Guideline for disinfection and sterilization in healthcare facilities, 2008. Available at: http://www.cdc.gov/hicpac/pdf/guidelines/Disinfection_Nov_2008.pdf [Accessed 5 May 2012]
6. Bond WW. Biological indicators for a liquid chemical sterilizer: a solution to the instrument reprocessing problem? Infect Control Hosp Epidemiol. 1993;14:309–312
7. Kohn WG, Collins AS, Cleveland JL, Harte JA, Eklund KJ, Malvitz DM. Guidelines for infection control in dental health-care settings – 2003. MMWR Recomm Rep. 2003;52(no. RR-17):1–67
8. Foster K Vast numbers of hepatitis C infections may come from routine dentistry. The Scotsman, 25 July 2001. Available at: http://www.rense.com/general12/mms.htm [Accessed 16 August 2012]
9. Gao G, Buskell Z, Seeff L, Tabor E. Drift in the hypervariable region of the hepatitis C virus during 27 years in two patients. J Med Virol. 2002;68:60–67
10. Rense J Invasive ‘sterilized’ re-usable medical/dental instruments as modes of CJD transmission, 2003. Available at: http://www.rense.com/general46/inva.html [Accessed 20 November 2010]
11. Piazza M, Borgia G, Picciotto L, Nappa S, Cicciarello S, Orlando R. Detection of hepatitis C virus-RNA by polymerase chain reaction in dental surgeries. J Med Virol. 1995;45:40–42
12. American National Standards Institute. Good hospital practice: steam sterilization and sterility assurance. ANSI/ AAMI ST46-1993. 1993 Arlington, VA Association for the Advancement of Medical Instrumentation
13. 29 CFR Part 1910.1030, occupational exposure to blood borne pathogens; final rule. Federal Register 56(235):64004-182, 1991
14. Foundations in continuing dental education. Available at: http://www.fda.gov/cdrh/ode/germlab.html [Accessed 11 October 2007]
15. Infection control in dentistry bda advice sheet A12. Available at: http://www.virox.com/msds/pdf/bda-cross-infection.pdf [Accessed 12 May 2012]
16. Infection Control Manual. Available at: http://www.dental.ohio.gov/icmanual.pdf [Accessed 12 May 2012]
17. Infection Control in the Health Care Setting. 2002 Canberra Australian Government Publishing Service
18. Summit JB, Robbins W, Schwartz RS Fundamentals of operative dentistry: a contemporary approach. 20012nd ed. Carol Stream, IL Quintessence Publishing Co. Inc.:345–351
19. Vickery K, Pajkos A, Cossart Y. Evaluation of the effectiveness of decontamination of dental syringes. Br Dent J. 2000;189:620–624
20. Smith A, Letters S, Mchugh S, Bagg J, Perrett D, Lange A. Residual protein levels on reprocessed dental instruments. J Hosp Infect. 2005;61:237–241
21. Miller CH, Patenik CJ Infection control and management of hazardous materials for the dental team. 1994 St Louis Mosby-Year Book:191–192
22. . Infection control recommendations for the dental office and the dental laboratory. J Am Dent Assoc. 1992;123(Suppl):S1–S8
Keywords:

dental instruments; hepatitis C virus; infection control; sterilization

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