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J Am Dent Assoc, Vol 137, No 3, 363-371.
© 2006 American Dental Association
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RESEARCH

JADA Continuing Education

Measuring the validity of two in-office water test kits



Joseph A. Bartoloni, DMD, MPH, Nuala B. Porteous, BDS, MPH and Lee Ann Zarzabal, BS, MS


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Background. The authors conducted a study to determine the validity of two commercially available in-office water test kits compared with a spread plate technique using the gold standard dehydrated culture medium R2A agar for monitoring the quality of dental treatment water.

Methods. Over a 12-week period, one author monitored nine dental units in a dental school that each were equipped with an independent water reservoir. The author collected 351 split samples, cultured them using three test methods, counted bacterial colonies manually and assessed validity using two cutoff values: ≤ 200 colony-forming units per milliliter (CFU/mL) (an American Dental Association goal) and ≤ 500 CFU/mL (a Centers for Disease Control and Prevention [CDC] recommendation and a U.S. Environmental Protection Agency [EPA] mandate).

Results. Of the 351 split samples processed, the in-office test kits’ accuracy ranged from 25 to 69 percent, according to the ADA and CDC/EPA recommendations, compared with the R2A agar.

Conclusions. Overall, the in-office test kits underestimated bacteria levels, producing inaccurate measurements of bacterial levels compared with the R2A agar.

Clinical Implications. The data suggest that use of the two in-office test kits could result in a lack of compliance, owing to underestimating bacterial contamination with recognized recommendations for dental unit waterline quality.

Key Words: Dental unit waterlines; biofilms; water quality; water microbiology; dental infection control

During the past two decades, infection control practices have become a significant part of dentistry. The basis of dental infection control is to create and maintain a safe clinical environment to eliminate or reduce disease transmission between patients and dental health care personnel (DHCP). An emerging issue in dental infection control has surfaced: reducing the exposure of patients and dental staff members to microbes in dental treatment water (nonsterile water used for dental therapeutic purposes).

It has long been recognized that dental treatment water delivered by dental unit waterlines (DUWLs) can be contaminated by microorganisms originating from the water supply.1–6 Studies have shown that dental treatment water can be contaminated with up to 1 million microorganisms per milliliter of water.3,7–9 DUWLs provide treatment water via a network of small-bore tubing to dental handpieces, air-water syringes and ultrasonic scalers. The water is used for irrigation, cooling of dental burs and oral rinsing. The microorganisms found in dental treatment water vary by the geographic location and include fungi, amoebas, protozoa, nematodes, and saprophytic and opportunistic gram-negative bacteria.7 A majority of the detected microbes are of low pathogenicity and have been described as opportunistic pathogens.10 Some studies have shown that DUWLs harbor small numbers of opportunistic pathogens responsible for respiratory disease, namely Pseudomonas aeruginosa,11,12 Legionella species9,13–16 and nontuberculous Mycobacterium species,17 which can infect both patients and DHCP. In the past decade, two studies have found significant levels of endotoxin derived from the cell walls of gram-negative bacteria in DUWLs.18,19 These endotoxins may be significant enough to cause a febrile reaction in healthy patients and are a theoretical risk factor for increasing inflammation unnecessarily during periodontal surgery.

Biofilms constitute a protected mode of growth that allows for survival in a hostile environment.

Dental units are either connected to municipal distribution systems for potable water or fitted with an independent water reservoir. Both systems are subject to contamination. Depending on infection control practices, waterborne, planktonic microorganisms flow through the dental tubing and can settle on the inner surface of the tubing, initiating a chain of events resulting in colonization, microcolony formation and eventually biofilm development.20 Biofilms are defined as matrix-enclosed bacterial populations adherent to one another and to surfaces or interfaces.21 The bacterial biofilms consist of microcolonies on a surface, and within these microcolonies the bacteria have developed into organized communities with functional heterogenicity. Biofilms constitute a protected mode of growth that allows for survival in a hostile environment. These structures have been shown to be 500 times more resistant to antibacterial agents than are isolated colonies, and they are the cause of many persistent and chronic bacterial infections in patients.22

Singh and colleagues23 found that biofilms in DUWLs harbor a vast diversity of viable organisms, including 55 cultivated biofilm isolates. Other studies have shown that biofilms are the primary source of contamination in dental treatment water.7,24,25

Although the majority of biofilm microbes originate in the public water supply and generally do not pose a risk of disease for healthy patients, patients with weakened immune systems may be prone to infection from these organisms.26 To date, no disease transmission arising from DUWL microbial contamination has been documented, but there is irrefutable scientific evidence that dental treatment water is of poor microbiological quality and often would fail to meet U.S. drinking water standards.27–29

The U.S. Environmental Protection Agency (EPA) is responsible for establishing national drinking water standards. This organization has mandated a standard of 500 colony-forming units per milliliter (CFU/mL) of noncoliform bacteria or less for potable water in community water systems.30 Advisory organizations and professional associations also have issued recommendations for dental treatment water. In 1993, the Centers for Disease Control and Prevention (CDC) recommended to dental offices that they install and maintain antiretraction valves on dental units to limit retraction of contaminated fluids from the operating environment, and that dental offices flush the units at the beginning of the day and between patients.31 The CDC also recommended using sterile irrigants for surgical procedures and published infection control guidelines in 2003 that recommended that microbial levels in coolant/irrigant water used for nonsurgical dental procedures should be as low as reasonably achievable, at a maximum 500 CFU/mL or less.32

The 1995 ADA Statement on Dental Unit Waterlines urged increased efforts by researchers and dental manufacturers to improve the design of dental equipment to reliably deliver dental treatment water with 200 CFU/mL of heterotrophic, mesophilic bacteria or less in unfiltered output water.33 This upper level was derived from the standard set by the Association for the Advancement of Medical Instrumentation for water quality in hemodialysis units.26 In addition, the Organization for Safety and Asepsis Procedures issued a position paper regarding DUWLs that identifies practical recommendations for clinicians.34 Recently, Pankhurst35 developed a risk assessment protocol to analyze the hazards from biofilm organisms colonizing DUWLs on the respiratory health of the dental team members and patients.

Methods to control or eliminate DUWL contamination have been evaluated. Available technologies include independent reservoirs, chemical treatment, sterile water delivery systems, filtration and a combination of these methods. When choosing one of these technologies to address DUWL contamination, it is imperative to monitor the results periodically to ensure that protocols are performed correctly and that the devices are working according to manufacturers’ instructions.32 DHCP should consult with the water treatment system’s manufacturer to determine the recommended frequency of monitoring.32 Noncompliance and technique errors are the most common reasons for clinical failure that can be identified by a monitoring procedure.36 DePaola and colleagues36 have suggested that test methods for monitoring should be consistent with Method 9215 in the Standard Methods for the Examination of Water and Wastewater.37 Monitoring can be accomplished using a microbiological laboratory or an in-office test kit designed to measure the quantity of heterotrophic bacteria.

Method 9215, also called the heterotrophic plate count (HPC), can be determined by pour-plate, spread-plate or membrane-filter methods. HPC provides an approximation of the number of viable bacteria, which may yield useful information about water quality.37 The most common culture media used to grow and measure heterotrophic bacteria from water samples include R2A Agar (Becton, Dickinson and Company, Franklin Lakes, N.J.), HPC agar and plate count agar. The R2A agar is not as nutrient-rich as HPC or plate count agar, but it is better-suited for enumerating bacteria that grow in low-nutrient environments such as drinking water.38 Reasoner and Geldreich39 found that the R2A agar yielded significantly higher bacterial counts than did plate count agar, and they recommended incubating the R2A agar at 28 C for five to seven days. Williams and colleagues40 showed that using a low-nutrient medium (for example, dilute peptone) with reduced incubation temperatures (25 or 30 C) recovered greater numbers of bacterial colony-forming units than did using an enriched medium (for example, blood agar or trypticase soy agar). van der Linde and colleagues41 showed that microbiological surveillance of hemodialysis fluids could be performed more precisely with the R2A agar combined with room temperature incubation for 10 days.

Comparing different DUWL studies has indicated that there are validity problems with the various test methods in terms of bacterial colony-forming unit counts.42–44 This can be due to using different water systems, different dental units,45 and different kinds of culture media and incubation conditions to determine bacterial load.46 Noce and colleagues47 found that time and temperature selected for plate incubation could affect bacterial counts dramatically and, therefore, study interpretations and conclusions concerning the clinical acceptability of water exiting DUWLs. In general, they found that longer incubation times and higher temperatures yielded significantly higher counts. Most heterotrophic bacteria thrive in an aerobic, nutrient-poor, room-temperature environment, and incubation time and temperature should reflect the normal environment of these predominantly slow-growing organisms.48 Most microbiologists believe that using a low-nutrient medium like the R2A agar with incubation temperatures of 20 to 25 C and an incubation period of at least seven days yields the highest total bacterial numbers in an evaluation of waterborne bacteria.47 The R2A agar is considered the gold standard for measuring heterotrophic bacteria in water.

The dental literature includes studies evaluating the validity of in-office water test kits. Karpay and colleagues48,49 have found that an in-office test kit (HPC Dental Sampler, Millipore, Billerica, Mass.) was accurate and correlated well with the R2A agar, while Smith and colleagues50 determined that this method resulted in an underestimation of bacteria in DUWLs compared with the R2A agar.

We conducted a study to determine the validity of two in-office water test kits compared with a spread plate technique using the R2A agar.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Over a 12-week period, the primary author (J.A.B.) monitored nine dental units located in a dental school during his residency in community dentistry. All of the dental units were equipped with independent reservoirs. The primary author assigned three dental units randomly to each of three treatment groups: use of a continuous iodine-based cleaner (units A, B and C), use of an intermittent iodine-based cleaner (units D, E and F) or a no-treatment control (units G, H and I). He used tap water from each dental operatory sink as the source water to fill the independent reservoirs of each unit. He used the following steps to collect the water samples:

– flushed the sink faucet for one minute before dispensing water into the independent water reservoir and attached the water reservoir to the dental unit;
– flushed all of the dental treatment water outlets for 30 seconds and cleaned them with alcohol pads;
– from each unit, collected pooled 20-mL water samples from five outlets (a high-speed hand-piece, a low-speed handpiece, two air-water syringes and an ultrasonic scaler);
– placed collected samples into a 100-mL sterile collection bottle containing sodium thiosulfate to neutralize any residual halogen present.

The resident took water samples at baseline and once per week for 12 weeks on Monday afternoons before intermittent chemical treatment of the three designated units was begun. All of the dental units were in use on Monday mornings.

Validity determines how well the test measures what it is supposed to measure and can be evaluated only if there is an accepted independent method for confirming the test results.

The resident enumerated the bacteria using two in-office test kits: HPC Dental Sampler and Clearline Water Test Kit (Kerr/Metrex, Orange, Calif.), the latter of which is no longer on the market, and a spread plate technique using the R2A agar.

First, he processed the water samples directly without dilution using the in-office test kits according to the methods described by the manufacturers. The configuration of the in-office test kits permitted the draw-through of 1 mL of sample to affix microorganisms to the filter surface for subsequent culturing and allowed for direct counts of bacteria after incubation.

The remainder of the water samples was immediately taken to the laboratory. The resident made 10-fold serial dilutions in phosphate buffer (10–1, 10–2, 10–3) and thoroughly mixed each for 15 seconds. For each of three platings, he plated one-tenth of a milliliter of each dilution on the R2A agar using the spread plate method. All of the samples were incubated at 22 to 28 C for seven days. The resident also randomly tested operatory sink tap water from all of the units weekly throughout the study period using the R2A agar.

After incubation, the resident counted the colonies manually using magnification. Weekly, he calculated the bacterial counts of each in-office kit and the mean of the triplicate platings and made dilution corrections for the R2A agar. He assessed validity by measuring the sensitivity, specificity, positive and negative predictive values, and accuracy of the two in-office test kits using two different cutoff values (≤ 200 CFU/mL and ≤ 500 CFU/mL) and comparing these values with those of the R2A agar.

Validity determines how well the test measures what it is supposed to measure and can be evaluated only if there is an accepted independent method for confirming the test results (for example, the R2A agar). Sensitivity is the proportion of samples with bacterial counts exceeding the cutoff values that are correctly identified by the test. Specificity is the proportion of samples with bacterial counts below the cutoff values that are identified correctly by the test. In our study, a more sensitive test would be more useful clinically to rule out contamination (that is, counts above the designated cutoff values); a highly specific test would be useful to confirm the presence of contamination. Positive predictive value is the probability that samples with a positive test result will have bacterial counts that exceed the cutoff values. Negative predictive value is the probability that samples with a negative test result will have bacterial counts below the cutoff values. The predictive values are affected by the prevalence of contamination (that is, the proportion of DUWLs that were contaminated at the time samples were taken). The higher the prevalence, the better the predictive value (that is, a positive test more likely indicates contamination and a negative test indicates levels below the cutoff values). Accuracy measures the degree of agreement between the test and the gold standard. In this study, accuracy determined how well the tests detected contaminated DUWLs.

Often, validation processes are evaluated by using a Pearson correlation coefficient. This coefficient, however, fails to detect any departure from the 45-degree line. We chose Lin’s concordance correlation coefficient51 over the Pearson correlation coefficient since it is a reproducibility index that evaluates the agreement between two readings (from the same sample) by measuring the variation from the 45-degree line through the origin (the concordance line).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 CONCLUSIONS
 REFERENCES
 
Ninety-three percent of the operatory sink tap water samples met EPA standards. The resident immediately checked the remaining 7 percent and found them to be within EPA guidelines. He collected and cultured 351 split samples (three methods each testing 117 samples) of dental treatment water using the three test methods. The contamination prevalence was 87 percent for the ADA cutoff value and 73 percent for the CDC/EPA cutoff value when we tested the water using R2A agar. Only 8 percent of the control units, 28 percent of the units cleaned with continuous iodine-based cleaner and 44 percent of the units cleaned with intermittent iodine-based cleaner met either cutoff value using the R2A agar (Tables 1Go, 2Go and 3Go [page 369]). This indicates that a majority of the DUWL samples exceeded the recommended levels of bacteria owing to the ineffectiveness of the cleaning methods used.


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  		TABLE 1 Microbial counts for units cleaned with a continuous iodine-based cleaner (CFU/mL*).

 

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  		TABLE 2 Microbial counts for units cleaned with an intermittent iodine-based cleaner (CFU/mL*).

 

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  		TABLE 3 Microbial counts for the control unit (CFU/mL*).

 
Validity measurements are displayed in Tables 4Go (page 369) and 5Go (page 370) for both the ADA goal and the CDC recommendation/EPA mandate, respectively. These values are assessments of the correctness of the test results. The results show that both in-office test kits had low sensitivity, high specificity, variable positive predictive value, low negative predictive value and widely ranging accuracy for both cutoff values. This indicates that both tests did poorly in identifying water samples with high bacterial levels but did relatively well in identifying water samples with low bacterial levels. Also, when both in-office tests indicated high bacterial levels, the R2A agar confirmed these high levels in a majority of the water samples; however, only 50 percent of the Clearline Water Test Kit samples agreed with the R2A agar for the CDC/EPA guideline. On the other hand, when both in-office tests indicated low bacterial levels, the results did not agree with those of the R2A agar a majority of the time. Overall, compared with the R2A agar, both in-office tests underestimated the microbial levels throughout the study period, leading to inaccurate measurements of bacterial levels.


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  		TABLE 4 Validity measurements at ≤ 200 CFU/mL* (ADA Statement on Dental Unit Waterlines{dagger}).

 

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  		TABLE 5 Validity measurements at ≤ 500 CFU/mL* (CDC{dagger} recommendation{ddagger}/EPA§ mandate).

 
HPC Dental Sampler samples displayed higher sensitivity, higher predictive values, equivalent specificity and increased accuracy compared with the Clearline Water Test Kit samples at both cutoff values. This means that the HPC Dental Sampler was more likely to detect contamination than was the Clearline Water Test Kit. The predictive values, however, more accurately reflected what DHCP need to know. For example, if a water sample has a positive test result, does that mean the DUWL is contaminated? And if a water sample has a negative test result, does that mean the DUWL has bacterial levels below the designated cutoff values? Both products produced a significant number of false-negative test results, signifying missed identification of contamination.

The choice of cutoff values relates to the importance of both false-positive and false-negative test results. Moving the cutoff value from ≤ 200 CFU/mL to ≤ 500 CFU/mL changed all the validity measurements resulting in decreased accuracy (that is, we identified more false-negatives).

We confirmed the validation results using Lin’s concordance correlation coefficient. Table 6Go (page 370) displays Lin’s concordance correlation coefficient and its 95 percent confidence interval for each of the two in-office test kits. These results confirmed that the two in-office test kits could not reproduce the R2A agar method, verifying that neither was reliable.


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  		TABLE 6 Lin’s concordance correlation coefficient (95 percent confidence interval).

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
CONCLUSIONS
 REFERENCES
 
With increased emphasis on water quality standards as part of improved infection control in dentistry, monitoring should become part of an established, effective quality assurance program. The monitoring of DUWLs should be practical, cost-effective, easy to implement or interpret, and fully integrated into dental offices’ quality assurance programs. When monitoring is administered properly, it should yield high-quality data. An effective program must control all factors from sample collection, processing, culturing and data reporting.

The methods for recovering microbes from DUWLs have changed over time. Methods have evolved from use of nutrient-rich media with short and high-temperature incubation periods to the use of nutrient-poor media with long and room-temperature incubation periods. Dental practices have two ways to monitor DUWLs—shipping samples to a microbiology laboratory or using an in-office test kit. The in-office test kits are designed to measure bacterial colonies from undiluted water samples with incubation at room temperature for seven days. These test kits seek to eliminate the need for an incubator and do not require special packaging, handling or shipment of samples to a laboratory, which can simplify procedures for dental offices.

Only two studies have been conducted comparing the HPC Dental Sampler to the R2A agar, and to date no validity studies regarding the Clearline Water Test Kits have been conducted.49,50 Karpay and colleagues49 showed that the results from HPC Dental Sampler agreed with those of the R2A agar 92.6 percent of the time. However, they found that the HPC Dental Sampler generally underestimated colony counts when compared with the R2A agar, and the results of their study confirmed the overall superiority of the R2A agar spread-plating techniques.

Our results differ significantly from those of Karpay and colleagues.49 In their study, they treated all units weekly with sodium hypochlorite 1:10. We can only surmise that the difference in results may be owed to the increased efficacy of the sodium hypochlorite compared with the iodine products tested in our study. This may have resulted in significantly reduced bacteria counts below the ADA cutoff value, increasing the accuracy compared with our study (that is, fewer false-positives or false-negatives were identified). Also, Smith and colleagues50 showed that some bacteria failed to grow on the HPC Dental Sampler compared with the R2A agar, resulting in reduced microbial counts, which was confirmed by our findings.

The validity results of our study showed that test measurement methods have a dramatic effect on the HPC values. Both the HPC Dental Sampler and the Clearline Water Test Kit consistently underestimated bacterial levels compared with the R2A agar, leading to inaccurate counts. The ingredients used in both in-office tests were proprietary. Our results suggest that those ingredients do not mimic those found in the R2A agar, which could have resulted in the discrepancies between test measurement methods. No specific medium, temperature or incubation time will allow for ideal conditions to measure all microbes at all times. As Karpay and colleagues48 concluded, each combination of microbes yields an estimate that varies as a result of the proportion of different species present and their characteristics.

Monitoring is becoming more widely accepted as a means of assessing compliance with methods used to control or eliminate DUWL contamination. It seems prudent to use a medium that will provide the highest estimate possible when documenting bacterial levels related to treatment protocols. Regardless of the treatment methods, monitoring should be done to ensure that protocols are followed. The development of an accurate in-office test kit that is inexpensive and simple to use would further encourage the adoption of a regular monitoring program for DUWLs.

Dentistry needs to adopt a standard protocol for the handling of water samples. The data from our study highlight the need for continual development of accurate in-office test kits that can provide valid bacterial counts. It would be advisable for dental manufacturers to develop a medium that is consistent with the R2A formulation. Without accurate in-office test kits, many dentists will be reluctant to monitor DUWLs using microbiological laboratory testing.

Researchers, manufacturers and dentists must share a common goal of improving the quality of dental treatment water. This will result in dental practices’ being better able to implement acceptable procedures when addressing this public health issue. It also will ensure that both patients and staff members are protected appropriately.


   CONCLUSIONS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
REFERENCES
 
There is minimal epidemiologic evidence that microbial contamination from DUWLs is a significant infection risk for patients and dental staff members. However, the potential for infection does exist, and the effects of DUWL contamination require further investigation. Therefore, every effort must be made to improve the microbiological quality of dental treatment water to meet recommended levels established by the ADA and CDC/EPA.

In our study, both in-office test kits demonstrated questionable validity, which underestimated the bacteria levels compared with the R2A agar. Use of these test kits in dental offices may result in a false sense of security and a lack of compliance with recognized recommendations for DUWL quality. Research is needed to develop standard ways to monitor bacteria levels accurately.


   FOOTNOTES
 

Dr. Bartoloni was a dental public health resident, Community Dentistry, University of Texas Health Science Center, San Antonio, when this study was conducted. He now is a dental public health consultant in the U.S. Air Force, Brooks City-Base, Texas. Address reprint requests to Dr. Bartoloni at 2509 Kennedy Circle, Brooks City-Base, Texas 78235-5116, e-mail "joseph.bartoloni{at}brooks.af.mil".


Dr. Porteous is an assistant professor, University of Texas Health Science Center, San Antonio.


Ms. Zarzabal is a statistical consultant, University of Texas Health Science Center, San Antonio.


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 CONCLUSIONS
 REFERENCES
 

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