Elsevier

Water Research

Volume 37, Issue 9, May 2003, Pages 2063-2072
Water Research

Precision and accuracy of an assay for detecting Ascaris eggs in various biosolid matrices

https://doi.org/10.1016/S0043-1354(02)00597-3Get rights and content

Abstract

This paper presents quality assurance data and quality control data on the recovery of Ascaris suum eggs from various biosolid matrices: acid-treated, alkaline-treated, amended-soil blended, and lagoon stored biosolids. Over a period of years, the same procedure, the “Tulane Method,” was performed on different matrices, and in this work, the data collected on the recovery of the eggs from the different matrices is examined and compared. The egg recoveries are discussed in terms of precision (the comparison of the recovery of eggs from a sample processed in duplicate) and in terms of accuracy (the percentage of eggs recovered from a sample to which eggs were added at the beginning of the extraction procedure). This form of quality analysis/control is typically called the “Split/Spike” method. This method of biosolid processing for helminth egg recovery had an overall accuracy of about 60% or greater and a percent variation from the mean density (an indirect method of assessing precision) of only 3–35%.

Introduction

The use of helminth eggs to monitor pathogen inactivation in biosolids is due to their higher survival rate (years) over bacteria (months), protozoa (weeks) and viruses (months) [1].

Over the last few decades, different procedures for detecting viable and non-viable helminth eggs in municipal wastewater biosolids have been developed. Unfortunately, no assay has yet been universally accepted for general use. One reason for this may be the lack of published QA/QC data on these assays. When our studies began in the 1970s, there were few published procedures for examining biosolids [2]. The few procedures that had been used by workers were mostly slight modifications of methods used to examine human and animal feces for parasites and were not suited for the examination of sample volumes greater than one to several grams. Because one needed to separate a small number of helminth eggs from a relatively large amount of solids, different procedures had to be developed for examining biosolids.

The two types of procedures most commonly used to recover helminth eggs from feces, biosolids, and soils were flotation and sedimentation. By their nature, sedimentation procedures separate the heavier particles, including helminth eggs, from the lighter particles in the sample. Consequently, the concentrated sediment recovered in a sedimentation procedure contains small, dense particles that concentrate in a manner similar to the helminths eggs and this makes the microscopic examination of the sediment difficult. When flotation procedures are used, many of these small particles, which are denser than Ascaris eggs, do not rise to the surface of the flotation solution and are thus eliminated [3]. Ultimately, through a series of experiments comparing flotation and sedimentation, it was found that flotation procedures worked better than sedimentation procedures for biosolids.

Flotation procedures utilize solutions of sucrose or metallic salts having a specific gravity greater than that of the helminth eggs in order to separate the eggs from the heavier particles in the biosolids. We tested sucrose and a variety of salt solutions including those of NaCl, CaCl2, NaNO3, MgSO4, ZnSO4 and HgI2. Sucrose was difficult to use in this procedure due to its thick, highly viscous nature requiring longer centrifugation times to obtain equivalent separation. Some of the salt solutions did not work well because they tended to precipitate when exposed to detergents during the process. MgSO4 and ZnSO4 were both found to work well, and a solution of either with a specific gravity of 1.20 was found to be suitable. For economic reasons, USP grade of MgSO4 was used for most of these studies; its cost was a fraction, about 1/10th of the cost of a reagent grade MgSO4 or ZnSO4.

The analysis procedure that was developed to detect eggs utilizes a number of steps prior to the flotation step. These include the initial dilution of the sample in water, homogenization by blending, and passage through one or more sieves to remove larger particles [3], [4], [5]. Then gravity sedimentation is used to separate the heavier particles from the unwanted small and less dense particles, such as bacteria, and to eliminate soluble material. An anionic detergent is added to the washed sediment to solubilize the organics and to aid in releasing eggs that may be adhering to larger particles. A variety of detergents were compared in our studies, including triton, Tween 80, and 7X®. We selected 7X® because better recoveries were obtained than when other detergents were used with MgSO4 or ZnSO4 and 7X® did not form precipitates when mixed with the salt solutions.

The recovery of helminth eggs from the surface of the flotation solution is usually done by one of two methods. In one, the upper layer of the flotation solution is decanted into a container, this solution is diluted with water to lower the specific gravity of the solution to below that of the eggs, and then the particles are collected in the sediment by gravity settling or centrifugation. In the other method, the flotation solution is poured through a sieve with a pore size small enough to retain the eggs. The eggs are then recovered by washing them off the sieve into a container. The latter method allows very small particles to be eliminated since they pass through the sieve and are discarded. This reduces the amount of particulate matter in the sediment that is to be examined microscopically. The latter method was chosen for our assay method.

The scope and aim of this paper is to review the quality assurance and control (QA/QC) data collected from our studies and to present the precision and accuracy of the Tulane assay with different types of biosolids. Over the past years, our team has worked on the recovery of helminth eggs from biosolids following different treatment processes. In this paper, efficiency and precision data on the assay we have developed are presented. Also, appropriate pretreatment and storage techniques for biosolids used in monitoring treatment process performance and analysis procedures are described.

Section snippets

Sources and recovery of eggs used in assays

Dr. Blair Leftwich in Lubbock, TX recovered the eggs used in all these assays from the feces of swine. The fecal droppings of pigs on a farm near Lubbock were collected by washing them from the floor of concrete holding pens and then placing the collected material in large containers to allow for settling. After 1 day, the supernatant was discarded and the sediment transported to the laboratory where it was passed through sieves (10 and 20 gauges) to remove the larger particles. The remaining

Synox-treated biosolids

The Synox process is an acid-treatment process (pH=3.0) with addition of sulfuric acid, ozone and nitrous acid. The quality-control data for the Synox experiments conducted in 1990 [4] are summarized in Table 1, and those that were conducted in 1991 [5] are in Table 2. The overall precision of the parasitologic analyses in 1990 was 9.7% (mean of the mean standard deviations) for all the samples examined irrespective to exposure time and percent solids. The precision of the sludge samples from

Discussion

Different workers have developed various assays for helminth eggs but no single assay is universally accepted. This is mainly due to the lack of published QA/QC data for the protocols used by different workers. J.C. Burnham showed a 10% recovery for Ascaris eggs in 1988 [13]. Huyard et al. [14] reported a 50% recovery of Ascaris eggs using a refined White House document procedure. They stated that this was a four-fold increase in recovery as compared to the procedure they had previously used.

Conclusions

The analysis of the assay reported on here for the analysis of the viability of Ascaris eggs from biosolids for PFRP or Class A equivalency testing revealed the following:

  • (1)

    The assay is fairly accurate for anoxic/acidic biosolids at 75–80% with a precision of around 10–15%.

  • (2)

    The assay's precision drops from 10% to 30% as the pH rises or soils are blended with biosolids.

  • (3)

    As the samples pH rises to around 12 (alkaline stabilization), the accuracy of the procedure decreases from 75% to 58%.

  • (4)

    Other assays

References (16)

  • National Research Council. Biosolids applied to land advancing standards and practices. Washington DC: The National...
  • Reimers RS, Little MD, Englande AJ, Leftwich DB, Bowman DB, Wilkinson RF. Parasites in southern sludges and...
  • Reimers RS, Little MD, Englande AJ, McDonell DB, Bowman DB, Hughes JM. Investigation of parasites in sludges and...
  • Sobsey MD, Hall RM, Buress AE, Blythe RD, Little MD, Reimers RS. Evaluation of the synox process for disinfection of...
  • Reimers RS, Little MD, Lopez A, Kitata KM. Final testing of the synox municipal sludge treatment process for PFRP...
  • Reimers RS, Little MD, Akers TG, Henriques WD, McDonell DB, Mbela KY. Persistence of pathogens in lagoon-stored sludge....
  • Leftwich DB, George DB, Reimers RS, Little MD, Klein NA. A field investigation of ascaris ova survival in domestic...
  • Little MD, Badeaux R, Reimers RS, Leftwich DB. Sensitivity of a procedure for the parasitologic examination of soils....
There are more references available in the full text version of this article.

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