In the Clinical Laboratory, we employ numerous methods to sample microorganisms from the skin of human volunteers. These sampling methods are not normally part of a microbiologist’s college curriculum, and newly hired technicians need to be trained appropriately to perform the methods in our laboratories. Two of the methods primarily employed at BioScience Laboratories, Inc. (BSLI) are the Glove-Juice Sampling Procedure and the Cylinder-Sampling Technique (also sometimes call the Cup-Scrub Method). In addition to proper training, we also employ statistical techniques to help identify those requiring additional training and for improving upon our methods.

For studies that involve evaluating Handwash products, we employ the Glove-Juice Sampling Procedure to sample microorganisms from the hands. A sterile glove is placed over the hand and a measured volume of sampling fluid is placed in the glove. The method of gloving can differ depending on the type of study. Hands may be wet, dry, held higher than elbow, or held lower than elbow. The glove is secured at the wrist using a tight-fitting elastic band. The hand is then massaged in a standardized manner for 1 minute by the technician. Following the massage, a sterile pipette is used to draw out a volume of fluid that is then diluted and plated. As you can see, the multitude of steps would have to be practiced and standardized so that all technicians perform the procedure in the same manner, and mistakes that may cause the loss of data are not made.

Sampling microorganisms from a small area of skin is performed using the Cylinder-Sampling Technique. A small (~1 in diameter), stainless steel cylinder is held tightly on the skin with one hand, while the other hand uses a pipette to place a small measured volume of fluid into the cylinder. A rubber-tipped glass rod is then rubbed against the skin inside the cylinder for 1 minute using a standardized, sweeping motion. A new pipette is used to transfer the fluid to a test tube. The procedure is repeated, with the second volume of fluid being removed and pooled with that fluid from the first sample. In addition to being trained to perform the procedure, technicians also need to learn the importance of cylinder placement and how to deal with sampling at a multitude of anatomical sites. This technique, too, requires lengthy training and practice.

Over my tenure at BSLI, Dr. Paulson has always employed statistical methods, such as Exploratory Data Analysis, to “look” at the data to determine and identify data points that are extreme “outliers” compared to the other data points. This statistical technique is used to identify human volunteers who may have not followed product restrictions or those who did not perform a wash procedure correctly, as well as help to determine if a noted incident that occurred had an affect on the data. EDA can also be used to evaluate the sampling methods and the technicians performing them… that is, identifying technicians who may require additional training. Conversely, EDA can also be used to identify technicians who perform the procedure well. The sampling method then can be adjusted to how they perform it, thereby decreasing variability and increasing the acuity of the conclusions drawn from the data.

Properly training technicians at BioScience Laboratories, Inc., is always a priority. We understand that our methods are non-traditional, but data generated by them is important to evaluating the true efficacy of a product. The initial training of a technician is vital to the conductance of the study, but continually monitoring the technicians’ performance and identifying minor adjustments has been and always will be a continual process here at BioScience Laboratories, Inc.

Christopher M. Beausoleil, CCRP

 Historically, the effectiveness of virus inactivation is estimated using procedures for detecting the loss of the virus’ ability to replicate. Until recently, this approach could not be applied to the majority of noncultivable viruses. There have been several attempts to develop indirect infectivity assays in order to solve the problem. One attempt is the enzymatic pretreatment of inactivated viruses with proteinase K and Rnase . Different research groups presented contradictory results on the reliability of this method. For the most part, it is because the effectiveness of the enzymatic pretreatment depends on the type of inactivation producing different levels of disruption, and hence, different levels of accessibility of peptide bonds to proteinase. Moreover, enzymatic pretreatment represents a secondary inactivation of presumably inactivated viruses, which might further enhance virus disruption.  Relatively reliable results were obtained by assessing loss of the inactivated virus’ ability to bind cellular receptors. The cell attachment capabilities of Poliovirus, Feline Calicivirus and HAV were lost in response to high temperature and hypochlorite treatment. However, UV inactivation did not prevent HAV from binding to the cellular receptor. This indicates that alterations of virus cell-binding sites are not the only mechanism of inactivation.   As a whole, detection of virus viability remains essential. The infectivity assay by means of cell culture has been the gold standard against which all new technologies for virus detection are evaluated. Methodologies combining cell culture and molecular techniques – for example, ICC-PCR (Integrated with Cell Culture PCR) and its modifications – have been used successfully for rapid detection of infectious enteroviruses and adenoviruses in environmental samples. The ICC-PCR technique is based on PCR analysis of the cell culture after incubation with test samples. Cell culture incubation allows elimination of non-infectious viruses and multiplication of infectious viruses, which then are detected by PCR. This method provides rapid and reliable detection of virus viability. Although ICC-PCR was developed originally for the detection of infectious viruses in environmental samples, the general principle of this method is applicable to detection of a replicating viruses in cells – for instance, the strand-specific PCR detection of Hepatitis C virus (HCV) in peripheral blood mononuclear cells. In this application, viral replication in cells is detected in terms of a HCV replicative intermediate, which is a negative-strand RNA. Detection on the basis of the negative strand means that false positive results due to amplification of RNAs of non-replicating viruses are avoided. Generally ICC-PCR is a very promising approach which with some modifications can be applied to identification of virus inactivation.

Volha Dzyakanava, PhD – Manager Virology Laboratory

To be successful in the marketplace eye care cosmetics must be non-irritating to the consumer.  Traditionally, a Draize rabbit eye test has been used to determine the safety of cosmetic products before sale to the general public.  The Draize Test involves applying the products directly to an animal’s eye.  The animals are observed for up to 14 days for signs of irritation (redness, swelling, discharge, cloudiness, or blindness in the tested eye).  Animal rights concerns, the cost of testing, and current European legislation banning cosmetics that have been tested using animals begs for an alternative model.  The MatTek  EpiOcular™ Tissue Model is a highly reproducible human cell-based in vitro tissue model that can be used to replace the traditional Draize ocular irritation testing.  The EpiOcular™ tissue model consists of normal, human-derived cells similar to cells found in the cornea (eye).  These cells are cultured on specially prepared cell culture forms creating multi-layered structures that are mitotically and metabolically active and induce the same inflammatory response as the human eye and can be used for detection of ocular irritation.  

We performed a collaborative study with MatTek to determine if the EpiOcular™ tissue model could also be used to differentiate between ultra-mild eye care cosmetic formulations.  Ultra-mild classifications can not be determined using the standard Draize rabbit eye test due to the insensitivity to the low levels of irritation induced by these products.  For the mildness testing, 10 commercially available mascara products, all with non-irritating or hypoallergenic claims, were purchased representing a broad range of manufacturers including Almay, Revlon, L’Oreal, Maybelline, and Covergirl.  The mascara products were tested using the EpiOcular™ model with an extended time exposure protocol (up to 24 hours).  The mascaras were applied to the surface of the ocular cells and remained in direct contact with the tissues for 8, 16, or 24 hours.  Following the exposure times, the mascaras were removed and the tissues and exposed to MTT for 3 hours.  Following the MTT exposure the tissues were removed from the MTT and rinsed with a Phosphate Buffered Saline solution. An alcohol extractant was added to the tissues following the MTT exposure.  The extraction process was allowed to proceed overnight.  The optical density (color change) of the extract was read the following day using a spectrophotometer.  If the mascara was non-irritating to the cells, the cells remained viable and were able to metabolize the MTT, thereby reducing the MTT and creating a visible color change from red/orange to blue/purple.  The absence of this darkening indicated that the mascara was irritating to the cells.  These optical density readings were used to determine the percent (%) viability of each tissue following exposure to each of the 10 mascara products.  An ET-50 (the time it takes for the viability of the tissues to decrease to 50%) irritation score was then assigned to each mascara.   The commercially available mascaras showed ET-50 scores ranging from 8.7 hours to > 20 hours.  These results relate to very mild products whose irritation potential would not have been detected with the standard animal model.  As such, the extended time exposure protocol appeared to be a facile, cost-effective means to screen ultra-mild eye care cosmetics. 

LIV GRAVING AND JESSICA MCDONNELL