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9.7. Parameter-Specific Sampling Procedures Sampling procedures for regulated and
unregu-lated contaminants are presented in this section by group for the most frequently collected parameters.
Sampling procedures for newly regulated conta-minants and other parameters of concern can be found in EPA approved methods
(http://water.epa.gov/scitech/drinkingwater/labcert /methods_index.cfm)
The parameters and method-specific QC are found in the approved drinking water methods.
This section provides common parameter-specific sampling procedures that meet federal require-ments. State and local requirements can be dif-ferent or more stringent; therefore, the sampler should always check state and local sampling requirements before the start of sampling.
9.7.1. Field Data
pH, temperature, chlorine residual, and turbidity measurements are conducted in the field at the start of sample collection.
9.7.1.1. Applicable Regulations
The SDWA requires EPA to institute rules and regulations to ensure that drinking water is safe to consume. The SWTR and subsequent amend-ments, were promulgated to minimize pathogen contamination in drinking water. These are discussed in Section 9.2. Disinfection and filtra-tion for surface water and groundwater under the influence of surface water is required.
Chlorine residual must be checked to determine preservation, holding time, and, in the case of bacteria testing, data interpretation. Chlorine residual is measured when collecting samples in the drinking water program as free chlorine resi-dual. Total chlorine residual is measured by water supply systems during treatment design, process control studies, or for distribution system problem solving.
MCL or Sampling Values. The regulator should specify the requirements for pH, temperature, and chlorine residual values for compliance. The turbidity after conventional or direct filtration
should be ≤ 0.3 nephelometric turbidity units (NTU) in 95% of all monthly samples collected.
At no time should the turbidity exceed 1.0 NTU.
Slow sand and diatomaceous earth filtration systems must achieve a filtered water turbidity level of ≤ 1 NTU in 95% of the measurements each month (limits can vary by state). At no time can the filtered water turbidity exceed 5 NTU.
9.7.1.2. Number and Frequency of Drinking Water Samples
The pH, temperature, chlorine residual, and tur-bidity samples should be representative of the system’s filtered water. These measurements should be collected once per day and at the start of each sample collection. Turbidity must be monitored every 4 hours on combined filter efflu-ents, although individual states can reduce this requirement to once-per-day for slow sand or alternative filtration devices. For individual filters, turbidity must be monitored every 15 minutes.
9.7.1.3. Sampling Locations
The regulator should specify where the pH, tem-perature, and chlorine residual samples must be taken for compliance. One turbidity sample should be taken at each entry point into the distri-bution system.
9.7.1.4. Analytical Methods
The free chlorine residual method uses the rea-gent diethylphenylenediamine (DPD) in a colori-metric test to determine the amount of free chlo-rine available for disinfection. Total chlochlo-rine indicates how much of the added chlorine reacted with the indigenous components in the sample and can be determined by adding small amounts of potassium iodide to the DPD solution and recording the difference. The reagent used must be clearly identified as measuring either free chlorine or total chlorine. The results reported must also indicate free or total chlorine measure-ments.
Nephelometery is a standard test method for turbidity. It is based on comparison of the inten-sity of light scattered by the contained static water sample under defined conditions to the intensity of light scattered by a reference standard
in the sample container. The higher the intensity of scattered light, the higher the turbidity of the sample.
9.7.1.5. Sample Volume
Sample volume depends on the method used for each parameter; refer to Chapter 13 for a discus-sion of pH, temperature, and chlorine residual field measurements. The turbidity volume col-lected should be sufficient to:
Ensure a representative sample
Allow for replicate analysis (if required)
Minimize waste disposal
Comply with the analytical method used.
9.7.1.6. Bottle Type
Bottle type depends on the method used for each parameter; refer to Appendix B for a discussion of pH, temperature, and chlorine residual field measurements. Samples for turbidity should be collected in plastic or glass bottles, as specified by the selected analytical method.
9.7.1.7. Sampling Procedures
The field testing procedures for pH, temperature, and chlorine measurements are presented in Chapter 13. Chlorine residual test kits, including instructions, are available for drinking water testing to measure free or total chlorine. An example of a chlorine residual method can be found in Chapter 13. Most kits use DPD powder, tablets, or drops of liquid placed in a glass tube containing the sample. A color change after adding the reagent indicates that free chlorine is present. In some kits, the color is measured against a field comparator or portable spectropho-tometer showing concentration in mg/L or ppm.
If a free residual chlorine level of 0.1 ppm is detected, this test will show as a trace of pink color. Trace colors are visible by looking down the tube and against a white background, which ensures no reflection from any red or pink sur-rounding surfaces. Interferences in the measure-ment of chlorine can cause variations to the color or affect the color formation. Any unusual color formations or colors must be noted for proper data interpretation. Note in the FLB/FN the results in mg/L, the date and time of the sampler,
the name of the sample, as well as any observa-tions made during the sampling or analysis. If chlorine is not detected in the test, write “not detected.”
A one-time demonstration of technician profi-ciency should be on file in the training records.
The one-time demonstration should include a standard curve, a low-level measurement stan-dard, and an annual performance evaluation sample for chlorine and pH.
For turbidity sampling, all bottles must be tho-roughly cleaned and rinsed with turbidity free water prior to sample collection. Turbidity sam-ples are collected typically during general tap sampling, source water sampling from surface water, and source water sampling from wells.
Refer to Sections 9.6.1, 9.6.4, and 9.6.5, respec-tively, for general sample collection procedures.
9.7.1.8. Preservation Requirements
There are no preservation requirements associated with sampling pH, temperature, or chlorine resi-dual. Ensure that samples for turbidity are cooled to ≤ 6°C.
9.7.1.9. Holding Time
Samples for pH, temperature, and chlorine resi-dual must be analyzed immediately, which gener-ally means within 15 minutes of sample collec-tion. Samples for turbidity should be analyzed as soon as possible after collection. If storage is required, samples maintained at ≤ 6°C can be held for up to 48 hours.
9.7.2. Microbiological
This section covers specific sampling require-ments for the following microbiologicals:
Coliforms
Cryptosporidium
Giardia.
9.7.2.1. Coliforms
Coliforms are a group of closely related bacteria that are generally harmless, but do contain some species that are human pathogens. They are natural and common inhabitants of the soil and ambient waters, such as lakes and rivers. Coli-forms can be inactivated by treatment or die off
naturally in a manner similar to most bacterial pathogens and many viral pathogens. Therefore, if coliforms are found in the distribution system, pathogens also could be present.
9.7.2.2. Applicable Regulations
The TCR introduced in Section 9.2 was estab-lished by the SDWA with sample requirements primarily based on the presence or absence of total coliform bacteria rather than on the number of coliforms detected in the samples. The TCR exists to protect public health by determining the presence of disease-causing pathogens in drinking water.
The TCR became effective December 31, 1990 and supersedes the old National Interim Primary Drinking Water Regulations (NIPDWR) for maximum microbiological contaminant levels (effective June 24, 1977). The rule requires that total coliform-positive samples be further tested for fecal coliform or Escherichia coli (E. coli) and that a set of repeat samples be collected for each total coliform-positive sample. When coli-forms are detected, additional routine samples must be collected the following month.
9.7.2.3. MCL
For systems collecting fewer than 40 samples per month, no more than one sample per month is allowed to be positive for total coliform. For systems collecting greater than 40 samples per month, no more than 5% of all monthly samples are allowed to be positive. If a system exceeds the monthly MCL, the public must be notified within 14 days per the requirements established in the Public Notification Rule. If an acute MCL violation occurs (i.e., when any fecal coliform is detected on a repeat sample) the public must be notified within 72 hours.
9.7.2.4. Number and Frequency of Samples The number of samples that must be taken monthly is based on the population served by the water system. Table 9-2 provides an abbreviated list of minimum monthly monitoring require-ments. Many community systems routinely sample more than the minimum to monitor the system’s status.
Table 9-2. Microbiological Monitoring Frequency for Routine Sampling in PWSs
Population Served Minimum Routine Samples/Month*
25 – 1,000 1
1,001 – 2,500 2
2,501 – 3,300 3
* The regulator can specify a sampling frequency of less than once per month for selected systems. A CWS serving more than 3,300 must refer to the CFR. Contact your state drinking water representative for details.
Compliance with the TCR is based on the pres-ence or abspres-ence of total coliform bacteria. If all routine samples test negative (absent) for the presence of total coliforms, no additional testing is required for that monitoring period, regardless of whether sampling is routinely conducted monthly, quarterly, or annually. However, the rule requires that total coliform-positive samples (presence) be further tested for fecal coliform or E. coli and that a set of three repeat samples be collected for each total coliform-positive sample monthly, quarterly, and annually. Systems that collect one routine sample or fewer per month must collect a fourth repeat sample. If total coliforms are detected in any routine or repeat sample, the sampler must collect at least five routine samples the next month. Table 9-3 pro-vides an abbreviated list of the required frequency for repeat sampling.
Table 9-3. Monitoring and Repeat Sampling Frequency After a Total Coliform-Positive
Routine Sample
All repeat samples must be collected within 24 hours of notification of the total coliform-positive result, unless the state waives this requirement.
Each set of repeat samples must include the fol-lowing:
One sample at the same tap as the original sample
One sample within five service connections upstream
One sample within five service connections downstream.
If a system has only one service connection, the state can authorize the system to collect the re-quired set of repeat samples over 4 days, rather than within 24 hours, after being notified of a total coliform-positive result. Per state discretion, such systems can collect a larger volume repeat sample (or samples) instead of the standard 100-milliliter container (e.g., a single 400-100-milliliter repeat sample or two 200-milliliter repeat sam-ples in one or more sample containers of any size as long as the total volume collected is at least 400 milliliters). If a system collects more than one routine sample per month, the total volume required would be 300 milliliters.
9.7.2.5. Sample Locations
Samples for the TCR are collected from the distribution system based on the number and frequency requirements.
9.7.2.6. Analytical Methods
The method used for fecal coliform and E. coli analysis depends on that used for the total
coli-form test and must be listed as approved for compliance monitoring. EPA Method 1604 is commonly used for total coliforms.
9.7.2.7. Sample Volume
The sample volume depends on the analytical method used. However, at a minimum, one 100-milliliter sample must be collected per test.
9.7.2.8. Bottle Type
Sample containers for collecting coliform sam-ples must be sterilized, hold at least 120 millili-ters (4 ounces) of water, and contain sodium thiosulfate if the sampled water is chlorinated.
Bottles can be plastic or glass. The laboratory normally supplies the container. Glass-stoppered bottles must be covered with aluminum foil or char-resistant paper prior to sterilization. The covering on the top ensures that the seal between the stopper and the bottle is not contaminated, thus minimizing the potential for contaminating the sample when it is opened. Some labs furnish a single-service, sterilized, polyethylene bag or bottle containing a sodium thiosulfate tablet. The laboratory should perform sterility checks (one per lot) prior to releasing the containers for col-lecting samples. The laboratory normally pro-vides instructions to go along with the sample containers it has supplied. Refer to those when provided.
9.7.2.9. Sampling Procedures
The laboratory that supplies the sampling con-tainers might provide instructions for the type of monitoring being performed. Be sure to refer to those instructions when provided. Total coliform samples are collected typically during general tap sampling, source water sampling from surface water, and source water sampling from wells.
Refer to Sections 9.6.1, 9.6.4, and 9.6.5, respec-tively, for general sample collection procedures.
9.7.2.10. Preservation Requirements
Coliform samples must be refrigerated or cooled to ≤10⁰Cimmediately after sampling.
9.7.2.11. Holding Time
The laboratory must receive and begin analyzing coliform samples within 30 hours of collection.
If the laboratory is nearby, refrigerate the samples
with ice packs and deliver them directly. Sam-ples analyzed by an off-site laboratory should be shipped on ice via overnight service on the day they were collected.
9.7.2.12. Cryptosporidium and Giardia Cryptosporidium is a genus of protozoan para-sites potentially found in water and other media.
The recent taxonomy of the genus Cryptospori-dium includes two species that use humans as hosts: C. hominis and C. parvum.
Giardia is a genus of protozoan parasites poten-tially found in water and other media. The recent taxonomy of the genus Giardia includes one species that uses humans as hosts: G. lamblia (also called G. intestinalis or G. duodenalis).
Applicable Regulations. Under the 1996 SDWA Amendments, EPA must supplement the existing 1989 SWTR with additional pathogen control requirements. Several additions to the rule have resulted in the Long-Term 2 Enhanced SWTR, which focuses on the control of Cryptosporidium and microbial pathogens.
MCLG. The Interim Enhanced Surface Water Treatment Rule (IESWTR) has an MCLG of zero or a 99% removal for systems that filter.
Number and Frequency of Drinking Water Samples. Monitoring of source water is required to determine the level of treatment for Cryptospo-ridium. Monthly samples are required for the first 2 years, with the level of Cryptosporidium deter-mining the level of any subsequent treatment.
Most systems will not require additional treat-ment. A 6-year follow-up sample is required to ensure continued compliance. Small systems can monitor for E. coli and will monitor for Cryptos-poridium only if the E. coli concentrations exceed a specified level.
Sampling Locations. Samples are collected from source water for distribution systems.
Analytical Methods. EPA Method 1623 can be used to measure both Cryptosporidium and Giar-dia. Prior to analysis, the water sample must be filtered and the extraneous materials retained on the filter. Qualitative analysis is performed by scanning each slide well for objects that meet the
size, shape, and fluorescence characteristics of Cryptosporidium and Giardia oocysts. Quantita-tive analysis is performed by counting the total number of objects on the slide confirmed as oocysts. Quality is assured through reproducible calibration and testing of the filtration, immuno-magnetic separation (IMS), staining, and micro-scopy systems.
Sample Volume. The sample volume depends on the analytical method used. Generally, 10-liter bulk samples are taken.
Bottle Type. Ten-liter collapsible containers generally are used for collecting Cryptosporidium and Giardia. These containers must be sterilized, and are normally supplied by the laboratory. The laboratory should perform sterility checks (one per lot) prior to releasing the containers for col-lecting samples. The laboratory typically pro-vides instructions to accompany the sample con-tainers. Refer to those when provided.
Sampling Procedures. The laboratory that supplies the sampling containers can provide instructions for the type of monitoring being performed. Be sure to refer to those instructions when provided. Cryptosporidium and Giardia samples are collected typically during general tap sampling, source water sampling from surface water, and source water sampling from wells.
Refer to Sections 9.6.1, 9.6.4, and 9.6.5, respec-tively, for general sample collection procedures.
Preservation Requirements. Ambient water samples are dynamic environments and, depend-ing on sample constituents and environmental conditions, Cryptosporidium and Giardia oocysts present in a sample can degrade, potentially biasing analytical results. Samples should be chilled to reduce biological activity between collection and analysis.
If samples are collected early in the day, they should be chilled by storing in a refrigerator between 1°C and 10°C or on ice in a cooler. If the sample is pre-iced before shipping, the ice should be replaced with fresh ice immediately before shipment. If samples are collected later in the day, these samples can be chilled overnight in a refrigerator between 1°C and 10°C. This