Efficacy of current and novel cleaning technologies (ProReveal) for assessing protein contamination on surgical instruments☆

Efficacy of current and novel cleaning technologies (ProReveal) for assessing protein contamination on surgical instruments☆

21

D. Perrett, N.K. Nayuni Barts and The London School of Medicine, London, United Kingdom

21.1 Introduction Even simply cutting through tissue using a scalpel or manipulating it with forceps deposits a considerable amount of tissue on any exposed surfaces (Fig. 21.1). Table 21.1 shows the amounts of tissue (wet weight) deposited on a scalpel following cutting through tissue. Obtaining similar data for more complex instruments is difficult as it is not as easy to determine the surface area. Brain appears to be particularly sticky—an observation confirmed by a forensic pathology colleague, Professor Peter Vanezis. Extrapolating these results to larger instruments, with significantly larger and more complex surfaces, will result in hundreds of milligrams of residual tissue material. Although some 60% of the deposit by weight is intracellular water with some whole blood, the dried material on surgical instruments must consist of inorganic and organic salts, lipids, nucleic acids, sugars, as well as proteins [1]. The majority of these biomolecules are soluble in water and washing the instrument with detergents removes most remaining residues from stainless steel (SS) surfaces. An exception to this statement is that some classes of proteins are particularly adherent to many surfaces. The percentage protein composition ranges from some 8% in plasma and brain to ca. 20% in liver. Most of these cellular components are readily soluble in water or easily removed with the detergents commonly used in sterile services departments (SSDs). Unfortunately the hydrophobic proteins or those with many exposed SH groups are more difficult to remove from surfaces, particularly stainless steel. Although it has been appreciated for some time that instruments would fail SSD visual checks if excessive amounts of tissue were observed, smaller amounts of tissue or protein smears can be routinely missed. Most proteins are colorless and invisible to the eye. There is little or no correlation between visual appearance and the ­measured

☆

This chapter is a reprint of the chapter originally published in the first edition of Decontamination in hospitals and healthcare.

Decontamination in Hospitals and Healthcare. https://doi.org/10.1016/B978-0-08-102565-9.00021-2 © 2020 Elsevier Ltd. All rights reserved.

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Fig. 21.1  Brain tissue deposits on simple surgical instruments postoperation.

Table 21.1  Weight of tissue and protein deposited on a scalpel blade after cutting through three different organs (n = 6 blades per tissue). Brain

Kidney

Liver

Wet

Dry

Wet

Dry

Wet

Dry

Mass of tissue (mg) Mass of tissue (μg/mm2)

48 ± 8 80 ± 14

12 ± 1 19 ± 2

12 ± 4 19 ± 6

6 ± 0.5 9.5 ± 0.8

4.5 ± 1.4 7.5 ± 2.3

3.0 ± 0.4 5.2 ± 0.7

Protein (μg/mm2)

8.0 ± 1.4

4.0 ± 2

1.5 ± 1

amounts of residual protein [2, 3]. Around 2005 the Department of Health (DH) ­sponsored studies by a number of independent research groups that clearly demonstrated that significant amounts of protein were present on surgical instrument sets that were about to be used in theater in a number of hospitals [2, 4, 5]. In fact one of the authors of this chapter was even able to weigh the amount of tissue that could be removed from some instruments processed in some hospitals on an analytical balance (Table 21.2) [5]! All the research groups employed different analytical techniques and the hospitals involved were from different NHS regions but all the studies gave similar protein levels. The findings reported by the groups [2, 4, 5] were a force behind the present drive to improve protein detection on instruments.

21.1.1 Proteins Proteins consist of chains of amino acids linked via the amine group (NH2) of one amino acid to the carboxylic acid group of another (COOH) forming a peptide bond. There are some 20 amino acids in proteins. Although there are no firm definitions,

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Table 21.2  A study to investigate residual protein on reusable instruments about to be employed in the operating theaters of five different hospitals (μg protein/instrument determined using o-phthaldialdehyde/N-acetyl-l-cysteine assay). Hospital A 0 8 12 16 48 56 60 86 260

Hospital B 0 0 2 10 18 42 138 302

Hospital C 0 0 3 5 8 8 13 30 53 168

Hospital D 10 18 35 41 93 93 173 213 432 488

Hospital E 1 6 12 14 94 94

proteins typically contain from about 20 to over 100 amino acids in their structures. Smaller amino acid chains are called peptides or polypeptides. Clearly the possible number of proteins that can be formed by combining the 20 amino acids is massive. Based on the overall charge on the protein it is possible to classify them into three groups: acidic, neutral (hydrophobic), and basic proteins. Proteins are also classified according to shape, namely globular, helical, and sheet. Proteins are additionally recognized by the sequence of amino acids in their polypeptide chain (primary structure), regular and recurring arrangement of primary chain, e.g., helix formation, gives rise to their secondary structure. Further structural changes, e.g., 3D bending and twisting of the chain into more compact structures, can be due to strong bonds forming between nearby sulfhydryl groups forming the tertiary structure. Finally the grouping of two or more polypeptide chains into a multisubunit protein held together by relatively weak bonds can occur, forming a quaternary structure. Hemoglobin is an example of such a complex protein with two pairs of alpha and beta globins encasing a central iron (heme) group.

21.1.2 The prion protein Naturally occurring prions (PrPc) are encoded by a gene on chromosome 20 and are active within the brain and other nerve tissues. PrPc is a protein composed of 253 amino acids glycosylated at positions 181 and 197. The exact role of PrPc is unknown but copper ion transport, cell signaling, protection, and synapse formation and even memory may be some of its functions. The protein-only hypothesis of transmissible spongiform encephalopathies (TSEs) suggests that this conversion of PrPc to the abnormal prion (PrPSc) is responsible for CJD. PrPSc is largely composed of beta sheets, which form strongly hydrophobic aggregates. Given the “sticky” nature of the aggregates, once in contact with instruments and surfaces they will no longer dissolve in water or even in the normal detergents used in SSDs. Prions can be transmitted via contact with infected tissue, body fluids, or contaminated surgical devices. The normal

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procedures employed in SSDs such as steam sterilizing do not render prions noninfective (for review see Ref. [6]). More information on prion proteins can be found in Chapter 14.

21.2 General principles of protein detection Although most proteins are colorless, many methods to measure proteins both qualitatively and quantitatively are available (for a review see Ref. [7]). The majority of these measurement methods rely upon functional groups within the structure of proteins reacting with chemical compound(s) forming colored or fluorescent derivatives. The majority of these reactions require the protein to be in free solution and the result is a measurement of the total amount of all proteins in the solution. For some proteins, specific assays based on enzyme reactions or immunoassays have been developed and are used in clinical diagnostics. The determination of protein residues on surfaces poses some difficulties as these proteins are bound to surfaces and are not necessarily able to react in the same way as they would in free solution. The methods that have been employed usually involved “removal” of either visible protein residues by swabbing or dissolving the invisible deposits using strong detergent solutions. The methods are summarized in Table 21.3. There is very little information on what particular proteins are found in the residual proteins. Recently Smith and Smith [8] studied the types of proteins released from dental extraction forceps when the working ends of forceps were boiled in 1% (v/v) sodium dodecyl sulfate (SDS). The proteins were separated using gel electrophoresis and followed by mass spectrometry. They identified 17 proteins including blood and bacterial proteins plus two unidentified proteins from decontaminated forceps. Most of the tests for total protein described in both the new guidelines, e.g., the Choice Framework for local Policy and Procedures (CFPP-01-01), and earlier guidelines, e.g., ISO/TS 15883-1, rely upon desorption of residual proteins from the instrument by wet swabbing techniques followed by a suitable chemical reaction to detect Table 21.3  Approaches to protein detection on instruments. Off instrument

On instrument

Others

Desorb (wipe) and test the swab for total protein Wash off with detergents and measure protein as amino acids following acid hydrolysis of washings Wash off with detergents and measure a specific protein in the washings Wash off with detergents and measure total protein in washings Cover or dip in a reagent and observe the color/fluorescence changes followed by recleaning “sandwich type” immunoassay for a specific protein, e.g., PrP Electron microscopy with X-ray scatter analysis Mass spectrometry Total organic carbon (TOC)

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the proteins desorbed on to the swab. Since the bulk of body proteins are hydrophilic, i.e., water soluble, they are readily removed by a simple water wash provided they have not been left to dry. Clearly the residual proteins after a hot detergent wash in an automatic washer-disinfector (AWD) are very unlikely to be easily removed by simple swabbing with cold water. The standard guidance to SSDs is to use rayon or cotton swabs wetted with water to “clean” a part of the instrument’s surface. The swab is then dipped into a visualizing reagent. Detection is either visual or by using simple instrumentation, such as a colorimeter or a reflectometer. The swabbing method was developed to assess the bacterial contamination of food utensils as long ago as 1917. Many subsequent researchers have shown that swabbing with water is very inefficient at removing spores, etc., attached to stainless steel. Variations in contaminant removal efficiency can be due not only to the type of materials involved and the nature of the cells and spores but also to the individuals doing the swabbing. The introduction to a recent paper by Rose et al. [9] reviews the background to the many early studies. Further they reported that recovery of Bacillus anthracis could be as low as 9.9% using rayon swabs rising to 43.6% if prewetted macrofoam swabs were used and following swabbing they were vortexed in liquid for 2 min. The same problems are also found with proteins. Nayuni et al. [10] found that water-wetted rayon swabs were very ineffective in removing proteins from SS. Instruments were contaminated with bovine serum albumin (BSA) or fibrinogen (a hydrophobic protein). After swabbing, chemical analysis found that 32 ± 4% and 61 ± 5% (n = 6) of the respectively proteins still adhered to the instruments. Addition of a detergent such as 0.5% Triton-X100 to the wetting solution only slightly enhanced desorption with 20.4 ± 3% BSA and 22.8 ± 2.8% fibrinogen (n = 6) remaining on the surface. Clearly the swabbing technique is not ideal for testing surgical instruments for residual proteins.

21.3 Current general methods of protein detection (ninhydrin, Biuret, dyes): Sensitivity, specificity, and validation A number of simple color tests for proteins were developed during the late 19th century but many require extreme conditions using strong acids and dangerous salts and are no longer suitable for routine use.

21.3.1 UV-Vis spectral measurements Since relatively few human proteins are colored, colorimetry and related color measurement techniques were restricted to proteins such as hemoglobin and whole blood. Obviously the standard laboratory practice of measuring the amount of protein in simple solutions by UV absorption at 280 nm cannot be applied to proteins bound to surfaces. The Soret spectral band of hemoglobin (414 nm) is used to detect hemoglobin directly but this cannot be easily applied to surface-bound material.

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21.3.2 Biuret and related methods It has been known for nearly two centuries that copper sulfate in alkaline solution turns from blue to violet/blue in the presence of proteins. This is called the Biuret reaction. By the 1880s, this was well established as a simple and reliable method to estimate protein concentrations in solution. An advantage of the Biuret reaction is that provided more than three amino acids are involved, it is relatively independent of protein composition since the complex forms with the peptide bonds. Unfortunately many compounds can interfere with its accuracy, yet today it continues to be one of the routine methods for protein estimation, and is employed in a number of both manual and automated assays for total plasma/serum proteins. Later Folin developed a versatile chemical reagent that was subsequently modified such that the new Folin-Ciocalteu reagent could be used to determine proteins. This reaction of copper with the tyrosine and/or tryptophan residues in proteins forms a cupric-peptide complex. In 1951, Lowry et al. [11] combined these established methods for determining protein by using Folin-Ciocalteu’s reagent to react with the cupric-peptide complexes formed in the classical Biuret reaction. Lowry’s method for protein determination results in a more prominent color change, i.e., from yellow to blue. Lowry’s method is one of the most highly citied methods in the scientific literature. A major improvement to Lowry’s method was published by Smith et al. [12] from the Pierce Chemical Company. They employed bicinchoninic acid (BCA), a specific reagent that chelates copper (Cu1+) ions forming a purple-blue complex (Plate VIII, between pages 358 and 359) that absorbs strongly at 565 nm. The reaction is faster (30 min) than the Lowry reaction and the sensitivity better at ca. 1 μg/mL. This chemistry is used in, for example, the Protest-Q kit (Table 21.4). However it is easy to generate both false negative and false positive results when used as a swab kit in SSDs. Recently Smith and Smith [8] employed the BCA assay to determine proteins desorbed from dental instruments. Biuret chemistry, or one of the above variants, forms the basis of a number of commercial decontamination test kits (Table 21.4).

21.3.3 Dye binding assays That proteins, in the form of natural fibers and skin, can strongly bind natural dyes has been known for eons. With the advent of synthetic dyes, the number of available dyes increased massively and some of these new dyes were shown to bind, albeit with varying specificity, to proteins and are routinely employed in histology. Such dyes have also been used to stain proteins separated by electrophoresis as well as to measure proteins in solution. The best known of these assays, the Bradford assay [13], uses the binding of the wool dye Coomassie Brilliant Blue (CBB) to proteins. The binding of CBB to a protein in an acidic solution changes the yellow/brown dye to bright blue (Plate IX, between pages 358 and 359). Since it is a binding assay the response is not linear with the amount of protein but the test is sensitive down to 100 μg/mL. Additionally, it exhibits differing response with different proteins. The simplicity of the assay has contributed to its continuing popularity in biochemistry. This assay combined with the swabbing approach is employed in at least two commercial protein test kits. A second dye binding assay from PEREG uses a pyrogallol red-molybate complex that has similar sensitivity to the Coomassie technique.

Chemistry

Mode

Trade name

Supplier

Address

Biuret

Copper binding

Pro-tect M

Medisafe

BCA—Lowry

Copper

Pro-Test-Q

Valisafe

Coomassie (Bradford)

Dye binding assay

Pyromol test

PEREG GmbH

Scope Check

Valisafe

DentaCheck

Valisafe

Clean-Trace surface protein plus Ninhydrin Protein Detection Test

3-M

Twyford Rd Bishop’s Stortford CM23 3LJ UK Twyford Rd Bishop’s Stortford CM23 3LJ UK Porschester 12 D-84478 Waldkraiburg Germany Twyford Rd Bishop’s Stortford CM23 3LJ UK Twyford Rd Bishop’s Stortford CM23 3LJ UK Carl-Shurz—Strasse 1 D41453 Neuss/Germany Waterside Rd Leicester LE5 1QZ UK

Ninhydrin

Ninhydrin

Albert Browne (Steris)

Sensitivity (μg/mL BSA) 1000 5 1000

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Table 21.4  Some commercially available test kits for residual proteins and their chemistries.

50

489

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21.3.4 Ninhydrin Ninhydrin is used in many bioanalytical techniques particularly for amino acid analysis method. Ninhydrin reacts with the α-amino group of primary amino acids producing “Ruhemann’s purple.” The chromophore formed is the same for all primary amino acids. The intensity of the color formed depends on the number and chemical nature of the amino groups being analyzed. The optimum pH for the overall reaction is 5.5. Ruhemann’s purple has a spectral maximum at 570 nm (see Ref. [14] for a review). Currently the ninhydrin test is employed by SSDs for residual protein detection on reusable surgical instruments as specified in CFPP-0101, the archived British Health Technical Memoranda (HTMs) 0101/HTM2030), as well as BS EN ISO 15883:2006Part 1). SSDs use rayon swabs wetted with water, to swab “cleaned” instrument surfaces before they are dipped in ninhydrin and heated to about 60°C for up to 1 h. If the swab turns purple then protein contamination is suggested, requiring the instrument to be rewashed. The positive control is a solution of the dibasic amino acid arginine. Such ninhydrin kits (see Table 21.4) are widely used. de Bruijn et al. [15] validated the ninhydrin swab test but with mg quantities of BSA while Lipscomb et al. [16] compared the ninhydrin assay to the Biuret test but used an unsuitable test soil containing free amino acids. A recent publication by the authors [10] has shown that the use of ninhydrin to test for proteins is extremely flawed. In laboratory studies, they showed that while ninhydrin detected amino acids with high sensitivity it was much less sensitive toward a variety of common proteins. This is because most proteins have only one free amino group at the N-terminal tail and although dibasic amino acids, i.e., lysine and arginine, occur in most proteins, steric hindrance limits the ability of ninhydrin to react with them. On a mass basis ninhydrin was some 40-fold less sensitive at detecting proteins than amino acids. Papers that recommend ninhydrin have usually employed whole blood or tissue homogenates and their authors have overlooked the fact that these biosamples contain large amounts of free amino acids as well as protein. A dibasic amino acid, arginine, is recommended in guidelines as the positive control and a solution is supplied with ninhydrin-based test kits. Arginine reacts readily with ninhydrin but it is not a protein and would be considered inappropriate as a control by most analysts. The combination of this lack of sensitivity with the poor desorption of proteins from SS means that the ninhydrin test generates a large number of false negative results from the decontamination process. It is concluded by the authors that this assay should no longer be employed to measure residual proteins on washed surgical instruments.

21.3.5 Spectrophotometric assay with o-phthaldialdehyde (OPA) derivatization Frister and Michels [17] introduced a method for measuring residual proteins based on an assay for amino acids that used o-phthaldialdehyde (OPA) in the presence of a mercaptoethanol to form an isoindole [18] (Fig. 21.2). They replaced mercaptoethanol with N,N-dimethyl-2-mercaptoethylammonium (DMMEA) and determined

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491 HO

O S H

OPA

O

+

HO

SH + R-NH2

Mercaptoethanol primary amine

N-R + 2H2O

Fluorescent isoindole

Fig. 21.2  The OPA reaction with mercaptoethanol and an amine.

the amount of detergent solubilized protein from instruments using a spectrophotometer set at 340 nm after a 2-min reaction time. This method is referenced in CFPP 01-01, the archived British HTMs 0101/HTM2030, as well as BS EN ISO 15883:2006-Part 1. McCormick et al. [19] using an experimental designs approach evaluated the spectrophotometric OPA assay for residual protein, with a test soil of blood, dehydrated hog mucin, and egg yolk placed on to microkeratomes. They concluded that this was an effective means for validating the cleaning of medical devices.

21.4 Methods of protein detection based on fluorescence There are many methods of protein detection that are based on fluorescence.

21.4.1 Basic principles of fluorescence The luminescences are physicochemical phenomena in which light is emitted from a sample. The luminescences include not just fluorescence, but also, among others, phosphorescence, bioluminescence, and chemiluminescence. Electrons excited by energy absorption must return to their ground state and for most molecules, this energy is released as heat. However in some molecules, some of the energy can leave as light. Fluorescence, the best known of the luminescences, is defined as the emission of electromagnetic radiation from suitable molecules, particularly in the visible region of the spectrum, after an initial absorption of a photon. Fluorescence is the result of a three-stage process. The excitation occurs when energy supplied by photon of light is absorbed by a fluorophore at a specific wavelength(s). The electron is forced into an excited state, which only exists for nanoseconds. During this time most of the energy is lost due to vibrational relaxation so depopulating the energy levels, eventually the electron must return to its ground state and in the process emit energy in the form of light (fluorescence). The light must leave at a higher wavelength (λEM), i.e., with lower energy than the excitation light (λEX). This change in wavelength is of the order of 50–100 nm and is called the “Stokes

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Table 21.5  Some common fluorescence derivatizing agents used with amino acids and proteins and their excitation and emission maxima. Compound(s)

Derivatizing agent

λEX

λEM

Primary amines

Fluorenylmethyloxycarbonyl (FMOC) o-Phthaldialdehyde (OPA) + thiol Dansyl chloride (DNS) Fluorescamine Fluorescein isothiocyanate (FITC) Acridinylmaleimide (AM) Monobromobimane (mBBr) 6-Aminoquinolyl-N-hydroxysuccinimidyl carbamate (AQC) o-Phthaldialdehyde (OPA) + thiol + detergent SYPRO—Ruby (SR) SYPRO—Orange (SO) Nano-Orange Cyanine dyes (e.g., Cy3) Fluorescein isothiocyanate (FITC)

265 340 372 390 493 360 394 250

315 425 500 475 518 450 490 395

355 450 472 485 488 493

455 618 570 590 509 518

Thioflavin T (Basic Yellow 1) (ThT) Congo Red

445 497

482 546

Free thiol groups General proteins

“Specific” proteins Amyloid Amyloid

shift” (see Table 21.5 for examples). Hence fluorescence detectors need two sets of optics—one set for excitation and the other set for the emitted light. Although if a very specific light source, e.g., a laser, is used the optics are simpler. A simple and common demonstration of fluorescence is achieved by observing a glass of tonic water in bright sunlight or under the “blue” lights commonly found in night clubs. The quinine in the tonic water fluoresces and the drink appears pale blue. When fluorescence is measured in solution, the fluorescence intensity [I] is related to the concentration of the fluorophore by Eq. (21.1): I ∝ I o cK

(21.1)

where I is the intensity of the fluorescence emitted, Io is the intensity of the incident light, c is the concentration of the molecule, and K is the system constant. Even in very fluorescent molecules the emitted light will probably not be more than 1% of the energy of the exciting light. Fluorescence instruments need to be designed to measure this small amount of light emitted by samples against a very dark background and, at the same time, exclude stray light from the source. From Eq. (21.1) it is also clear that sensitivity is dependent on the intensity of the light hitting the sample in the first instance plus the efficiency of the collection optics (K). Therefore the lamps used must have very high energies at the required excitation wavelength of the compound of interest to give the highest sensitivity. Typical high-­ energy lamps are xenon lamps (that have a continuous output from 240 to 700 nm)

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and mercury discharge lamps (discrete lines at 254 nm, 360 nm). Xenon lamps are very expensive while mercury lamps are very cheap (less than £20). Increasingly, lasers are found in fluorescence devices due to their very high energies and spot coverage. Unfortunately lasers only emit at specific wavelengths, which may not match the spectral characteristics of the fluorophores of interest. In fluorescence the excitation and emission wavelengths are selected by appropriate filters. The collection optics needs to gather the emitted fluorescence both selectively and with high efficiency and importantly minimize collection of scattered light. They usually use colored glass or high-transmission interference filters although monochromators can be employed. Fluorescence is usually of low intensity so must be measured by high-sensitivity devices such as photomultiplier tubes or charged coupled devices (CCDs) as found in digital cameras. When the instruments are suitably configured fluorescence is both highly sensitive and selective.

21.4.2 Native protein fluorescence Few compounds are naturally fluorescent but all proteins contain the amino acid tryptophan, which is fluorescent (λEX = 280 nm and λEM = 355 nm). However, 280 nm is only obtainable from only some lamps and being a UV wavelength it is damaging in use and needs quartz optics. Although UV-emitting lasers are available they are not suitable for routine use. So at this time detection of proteins on surfaces using their native fluorescence is not a viable prospect.

21.4.3 Protein derivatization to create fluorophores Nonfluorescent compounds may be reacted with specific chemical reagents to give fluorescent derivatives that can use more common light sources. Proteins too can be derivatized with a variety of reagents. Derivatization usually takes place prior to detection. So, ideally the derivatives should be stable: the derivatizing reagent should not in itself be fluorescent and react rapidly at room temperature. Possible reactive groups in proteins are the N-terminal amino groups, the epsilon amino groups of lysine and arginine, and any free thiol (SH) groups. Examples of some common derivatizing reagents for amino acids and proteins are given in Table  21.5. There are a rapidly growing number of fluorescent probes for proteins; see review by Sun et  al. [20]. For the most comprehensive listing see the website (www.invitrogen.com) or Life Technologies Corporation [21].

21.4.4 Fluorescent derivatization to determine residual proteins A number of groups (e.g., Refs. [4, 22]) have employed a technique of extensive removal of residual proteins using strong detergents followed by the quantitation of the desorbed proteins. Nearly all the published methods have used OPA in conjunction with various thiols. OPA reacts with the terminal and side change amino groups of proteins and a thiol compound to give a fluorescent isoindole (Fig. 21.3). This reaction was discovered in 1971 by Marc Roth, who used mercaptoethanol (ME) as the thiol.

494

Decontamination in Hospitals and Healthcare NH2

–S–S– O H H

–S–S– +

+

O OPA

O

NH2

HS

OH O

HN

N-acetyl-L-cysteine

NH2 Intact protein

NH2 + DTT + Triton-X100

O OH O S N H N O OH O

S SN H N

SH

O OH O S N H N

SH

SH

O OH O S N Denatured protein with fluorescent isoindoles H N

Fig. 21.3  The modified OPA reaction with N-acetyl-cysteine and a protein.

OPA/ME continues to be a favored reagent for amino acid analysis since a different isoindole is formed with each amino acid. No individual chemicals in the reagent possess native fluorescence so only the formed isoindoles are highly fluorescent. Major disadvantages of OPA/ME are the loss of sensitivity of the reagent over just a few days, the variable instability of the formed isoindoles, and the very unpleasant smell of ME. Use of automated derivatization for amino acid analysis has minimized the instability problems. Subsequently the lifetimes of both the reagent and isoindoles have been extended using other thiols (for a review see Ref. [23]). For residual protein detection, Verjat et al. [22] employed OPA with DMMEA to measure proteins released by various agents from instrument surfaces. The proteins were first acid-hydrolyzed for 24 h to release amino acids before assay with OPA/thiol. In order that proteins can react readily with OPA/thiol, in 2001, Perrett [published in Smith et  al. (2005) [3]] modified the reagent to include a detergent to denature the proteins prior to reaction. In addition, N-acetyl-cysteine (NAC), a more stable and much less smelly thiol, was used. This reagent was subsequently employed in a number of studies on decontamination [3, 5, 24]. The proteins were first desorbed by sequential sonication in a fixed volume of a strong detergent (Decon-90) and the concentration of protein in the combined washings was measured using OPA/NAC [3, 5, 24]. Zhu et  al. [25] adapted the reagent to work in a 96-well format with a

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p­ remade OPA ­solution from Pierce Chemical Co. Smith and Smith [8] used this format after desorbing proteins with an SDS solution and used DMMEA as a thiol.

21.4.5 In situ detection of proteins The semi-quantitative determination of proteins using fluorescent dyes has been a common technique in histochemistry and related areas for many years. When the fluorescent dye is coupled to a suitable antibody the technique is not only highly sensitive but can also be very specific. Cells or tissues stained with fluorescent probes are viewed using a fluorescence microscope in which the excitation source is focused directly onto the sample. The emitted fluorescence is viewed either directly or via a camera after being selected by an appropriate filter. The microscope system is most often configured such that the excitation light of the selected wavelength is reflected via a dichroic mirror and then focused on the specimen through the objective lens. Intense lamps, e.g., xenon, are therefore needed. The fluorescence emitted by the specimen is focused, by the same objective, and the mirror that is used for the excitation onto the detector. Since the majority of the excitation light is transmitted through the specimen, the background is much reduced, so an epifluorescence configuration gives a high ­signal-to-noise ratio along with excellent selectivity. The sensitivity of such an approach has been enhanced by using laser light sources.

21.4.6 Microscope-based systems Keevil and coworkers at the University of Southampton have been using episcopic differential interference contrast (EDIC) microscopy coupled with epifluorescence microscopy (EDIC/EF) to detect proteins for many years (Plate X, between pages 358 and 359) [26]. When used with a microscope objective offering an enhanced depth of field EDIC/EF can be used to examine proteins on surgical instruments that had been stained with suitable dyes. Lipscomb et al. [26] employed the technique to detect total protein on instruments that had been stained with Sypro Ruby (SR). SR and Sypro Orange were developed in 1996 by Steinberg et  al. [27] working for the company Molecular Probes as fluorescent stains for proteins separated by 2D gel electrophoresis. They bind noncovalently to most proteins and some lipopolysaccharides. Binding requires approximately 1 h. The fluorescent signals from a number of separate parts of stained instruments are captured by the EDIC/EF system and image analysis software is used to develop an image of the visualized area (Plate XI, between pages 358 and 359). The sensitivity is some 1000 better than the Biuret approach with a limit of detection of 175 pg/mm2. Later the system was adapted to detect amyloid protein using Thiazole T (ThT) dye [28]. With this stain a limit of 100 fg of amyloid per 1 μm diameter plaque was reported. However ThT dyes cannot discriminate between plaques composed of PrPsc and those composed of other amyloidogenic fibrils such as Aβ, a major component of Alzheimer’s disease pathology. Recently the same group [29, 30] developed a dual staining approach in which SR and ThT are used together so that total protein and amyloid protein could be detected at the same time in the same sample.

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21.4.7  X-Y scanning systems Some commercial spectrofluorimeters have the ability to use fiber optic cabling for excitation and detection to scan 2D surfaces such as chromatography sheets. Perrett demonstrated in 2001 that such an instrument could detect proteins reacted with OPA/ NAC on metal sheets with reasonable efficiency. However, the fiber optic heads could not track over the varying surfaces seen on surgical instruments without damage. The same approach is utilized in a system, called EFSCAN, developed at the University of Edinburgh by Baxter et al. [31]. Surgical instruments are dipped into a solution of fluorescein isothiocyanate (FITC) in dimethylsulfoxide. FITC reacts not only with the amino groups of proteins but also amino acids and other amines. The FITC solution is allowed to react for 15 min with proteins adsorbed to surfaces creating the fluorescent FITC derivative of the protein. Since FITC is itself fluorescent the specimen must then be washed with water to remove excess reagent and lower the background fluorescence. The FITC labeled proteins are then detected by the EFSCAN system, which consists of two fiber optic bundles attached to an epifluorescent head. Excitation is achieved using a 468 nm light-emitting diode (LED) and the emitted light is collected by the second fiber optic bundle and passed to a photomultiplier. The section of instrument to be measured is positioned via a computer-controlled X-Y stage below the fixed. Successive 0.5 mm2 sections can be scanned and a full image of the instrument generated via appropriate software. The sensitivity of the system is ng protein/mm2. This system, described in detail in Chapter 22, is being developed for commercialization.

21.4.8 Direct imaging systems In 2008 the OPA/NAC reagent of Perrett described above was modified so that it could be sprayed onto a surface contaminated with proteins. After drying, the surface could be illuminated with a suitable mercury lamp and the fluorescence be observed through a suitable glass filter. This approach requires the use of a dark room. Many gel-documentation systems used in protein and DNA research use an approach for detecting suitable fluorophores but in a compact laboratory instrument. It was envisaged that such an instrument, if equipped with a suitable mercury lamp and appropriate filter in front of a sensitive CCD camera, could be used not only to detect proteins on instrument surfaces but also to quantify the amount of protein by using appropriate software. Initially a system for 2D protein gel visualization from Syngene, Cambridge, the United Kingdom, was employed. First a digital black and white image of the sprayed instrument was captured and then the fluorescent image was captured. The two images could be overlaid using false color images to reveal where any residual proteins were. The color could be customized such that the intensity of the red indicates the intensity of fluorescence, i.e., amount of protein. By including spots of known amounts of protein in the exported fluorescent image it was possible to determine the volume of both the standard protein spots and the residual protein spots in order to quantify the amount of residual protein. All this was done manually using the Syngene’s image capture software, exported into D-Plot image analysis software to determine spot

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volumes with final calculations being done in MS-Excel. Typical images obtained are shown in Plate XI (between pages 358 and 359). At present these systems are only described in two patents [32, 33]. The system can visualize the entire surface of any surgical instrument that can be fitted onto a sheet of A4 paper. The detector response is linear in the range 0–50 μg spots for the calibration standard of BSA. The sensitivity of the system expressed as the smallest amount of BSA in a single spot is 50 ng. Working together the teams from Barts Medical School in London and Syngene in Cambridge have developed an automated system in a single unit designed specifically for SSD use. The system (i.e., spray plus detector), now called ProReveal, has been commercially available from early 2013 (Fig. 21.4). Users place a reprocessed surgical instrument in the tray of the ProReveal imaging system and then lightly spray with the OPA/NAC reagent. Images are captured after closing the instrument drawer and at the touch of an on-screen button, the system automatically displays an image of the instrument with any contaminating proteins on the visible surface (Plate XII, between pages 358 and 359). The built-in software indicates via an on-screen green tick or a red cross if this is a pass or fail of the decontamination process (seen in Fig. 21.4). Additionally, the software can be configured to show a value for the amount of residual protein found on the instrument. The process, taking less than 3 min in total, enables those in SSDs to rapidly perform sensitive in situ detection of proteins on reprocessed surgical instruments.

Fig. 21.4  The ProReveal system showing the spray and the imaging system.

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21.5 Other possible technologies Of course, scanning electron microscopy (SEM) can be used to image any deposits on surgical instruments. To identify protein in the deposits the SEM needs to be energy dispersive X-ray (EDX) spectroscopic analysis that can identify the elements, e.g., nitrogen in proteinaceous deposits. It is a method extensively employed in surface and material analysis. The method was employed by Chris Lowe’s group in the Institute of Biotechnology Cambridge, the United Kingdom, in their unpublished studies performed ca. 2004 on decontamination. More recently, it has been used by Baxter’s group in Edinburgh (2009). However, the SEM-EDX method is limited in its usefulness since it is not quantitative, gives only elemental compositions, and only small objects can be studied. Additionally, it requires an expensive SEM and using it is labor-intensive.

21.5.1 Mass spectrometry (MS) Recently, a number of mass spectrometry (MS) methods for surface analysis have become available. Perrett used matrix-assisted laser desorption/ionization time-of-flight (MALDI-ToF) MS to study the removal of the major plasma proteins from SS targets but clearances in the MALDI employed did not permit studies on anything thicker than a scalpel blade. In 2004, Graham Cooks at Purdue University developed a new desorption electrospray ionization (DESI) technique to analyze and identify organic compounds such as explosives on surfaces under ambient conditions [34]. An electrostatically charged water jet is directed on to a surface in order to desorb any analytes before they enter a standard quadrupole mass spectrometer. Unfortunately it requires the analytes to be a nonconducting surface so making DESI analysis of stainless steel surgical instrument is impossible. In 2007, Hiden Analytical (Cheshire, the United Kingdom) introduced plasma-assisted desorption/ionization mass s­pectrometry (PADI-MS) for surface analysis that does not require any preparation which can be used on metal surfaces and under ambient conditions. PADI is achieved by directing nonthermal atmospheric radiofrequency plasma onto the surface of interest. Desorption occurs from the surface and the ionized products are detected in real time by using an atmospheric sampling quadrupole MS. While it is unlikely that such MS techniques could be used routinely to monitor surgical instruments PADI might prove a useful technique to ascertain precisely what proteins remain following washing.

21.6 Strengths and weaknesses of new technologies All the new fluorescence methods described above are orders of magnitude more sensitive than the currently used “desorb and test” methods especially when it is considered that little residual protein is removed in the first place by swabbing. The new methods do, though, require investment in capital equipment ranging from about £10K to maybe £60K, depending on the system chosen. Running costs for consumables should be very similar per instrument tested to those incurred in buying the presently available tests

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such as the ninhydrin kit. Staff will also need to be trained not only in the operation of the equipment but also in the interpretation of the results. However these costs are small compared with the closure of whole operating suites due to failure of the SSD processes. All methods for measuring proteins need to be calibrated against an appropriate protein standard. However it must be pointed out that OPA/NAC, FITC, and SR can give differing responses to differing pure proteins. Responses can vary by a factor of 10. There are at least 500,000 different proteins and protein variants in human plasma but albumin is the dominant blood protein at ca. 55% and globins forming ca. 38%. Since the reagents react with the protein’s chemistry variations also occur, e.g., changes in room temperature or reaction times. Nearly all methods for measuring total protein use BSA as the calibrant since it is readily available and relatively inexpensive. The new fluorescent methods are no exception and are calibrated against BSA standards, so technically all results should be expressed as BSA protein equivalents although this is rarely done. Another problem is the units of measurement. Whereas the desorb/detect tests were simply a pass-fail system the new methods can generate what can, at least, be considered semiquantitative data. At present the three main research groups have generated data in different formats, for example, μg/total instrument, ng/examined spot, μg/m2, and ng/examined field. Although working with the same information the displayed data are calculated differently by the systems. All the systems can only quantify the amount of protein on one section of one side of an instrument: of course the instrument can be turned over and the measurement repeated on the underside. Statistically it is better to measure two instruments rather than two sides of the same instrument. Since the ProReveal system uses edge detection software to outline the 2D shape of the instrument first of all, it is then possible to calculate not only total protein per instrument side but also protein density, e.g., ng/mm2 for the whole instrument as well as calculate protein concentrations in hot spots. This allows an SSD operator to compare their processes across instruments of widely differing sizes, e.g., neurosurgical and orthopedic sets while generating data in a format similar to the other systems. Although individual SSDs may prefer differing units, it is expected that guidance on a standard unit may come from the Department of Health (England). At the time of writing no single standard exists for “acceptable protein levels” on reprocessed instruments. The BS EN ISO-15883 part 1 defines an acceptable level as below the limit of detection for any one of the three desorption and detection protein assays described previously. Confusingly the three methods report in different units! It states these to be 2 mg/m2 for the ninhydrin assay, 30–50 μg for the BCA assay, and 0.003 μmol for the spectroscopic OPA assay. Quoting three values is itself very confusing as well as the problems with the assays described in Section 21.3. It is expected that DH (England) will shortly announce the results of a statistical evaluation of protein levels on actual instrument as well as suggesting minimum testing frequencies. As with all new techniques there are some limitations and difficulties to the fluorescence methods of detection. Fluorescence is very sensitive, so care is needed not to contaminate instruments with either protein or reagents prior to measurement. The simplest manipulation can deposit sufficient protein from fingers, saliva, sweat, or the scalp on to the test piece to be measured. Depending on the excitation and emission

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wavelengths selected, chemicals in the environment might also be seen in images. For example, the wavelengths used with OPA/NAC also show the optical brighteners added to domestic washing powders and to whiten paper. So care is essential to avoid dust from paper and the black paper on which the instruments is placed in the viewer must be carefully chosen since most black paper is simply recycled white paper dyed black! Too much exposure to intense light sources can lead to photo-bleaching, i.e., loss of fluorescence. Scattered light, especially Tyndall scattering from the irregular surfaces of instruments as well as Rayleigh scattering, should be minimized in all systems in order not to confuse scattered light with fluorescence. The use of reagents that are nonfluorescent in themselves, e.g., OPA/NAC, and those with larger Stokes shifts are to be preferred. A concern, flagged in CFPP 01-01, is the safety of these fluorescent reagents, since unlike the swabbing methods, there is a small possibility of test chemical remaining on tested instruments. Toxicological data on the individual chemicals used suggests little concern at the low concentrations used to either manufacture the tests or in the final reagent used by SSD staff or the environment. All the reagents employed to date are readily removed by a second wash in an AWD. In addition, a second cycle in a fully validated AWD with standard detergents will remove any fluorescent proteins to below the limit of detection. However, there is still a very small possibility of traces of a reacted protein remaining on the instrument when reused. Repeated exposure to such nonself-proteins can theoretically induce an immunological reaction. Whatever is the amount of the protein, it will be much lower than the exposure to nonself-­residual proteins derived from previous patients on nonclean instruments. Guidance on the toxicological testing of other chemicals, e.g., both alkaline and enzymatic detergents and human proteins “damaged” by them by the regulatory agencies, is far from clear and some manufacturers do not appear to perform such tests.

21.7 Conclusion The major failings in the current recommended test procedures for residual proteins are overcome with the high-sensitivity fluorescent in situ tests now being developed and marketed. Such tests will not only improve quality control in SSDs but will also be of importance in testing the efficiency of available detergents and the development of both new detergents and improved AWDs. Of course a major failing of both the old tests and the new fluorescent ones is that they cannot directly determine protein residues inside endoscopes and more novel ideas will be required to do that.

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