Ann owup. tiyq.. Vol. 39, No. 2, pp. 223-233, 1995 Elsevier Science Ltd

British Occupational Hygiene Society Printed in Great Britam

ooO34878/95 S9.50+0.00

0003-4878(94)00116-2

THE MEASUREMENT OF AIRBORNE GLUTARALDEHYDE BY HIGH-PERFORMANCE LIQUID CHROMATOGRAPHY

John Cuthbert and John Groves Occupational Medicine and Hygiene Laboratory, Health and Safety Executive, Broad Lane,

Sheffield S3 7HQ, U.K.

(Receioed injnalform 22 September 1994)

Abstract-A published method for measuring personal exposure to glutaraldehyde has been evaluated. A change to the composition of the eluent used in the high-performance liquid chromatography analysis is suggested along with a more robust calibration procedure. Test atmospheres of glutaraldehyde were used to show that sampling efficiency is unaffected by humidity changes in the range 2&70% using sampling flow rates of up to 500 ml min- ‘. Samples can be stored at room temperature for up to 4 weeks after collection. The method has a detection limit of about 3 ppb for a 2 1. sample equivalent to about 1/65th of the current occupational exposure limit for glutaraldehyde.

INTRODUCTION

Glutaraldehyde [OCH-(CH,),-CHO] is widely used as an antimicrobial agent. It is irritating to the eyes, skin and lungs and has been assigned an occupational exposure standard (OES) in the U.K. of 0.2 ppm (0.7 mg md3) measured over a 15-min period. Exposure to glutaraldehyde has been measured in the past with a general calorimetric method for aldehydes using 3-methyl-2-benzothiazolinone hydrazone (MBTH). This method is, however, non-selective and will respond to other aldehydes such as formaldehyde. It is inconvenient as the air sample is collected into a liquid contained in a glass impinger, and for most airborne samples, it has to operate close to its detection limit of 20 ppb (assuming a sample volume of 11.25 l., i.e. the recommended flow rate, and 15 min sampling time). Thus a more selective and sensitive method for the measurement of personal exposure to glutaraldehyde was required.

The two methods available at the time this study began were those published by the National Institute for Occupational Safety and Health (NTOSH, 1989) and the Occupational Safety and Health Administration (OSHA, 1987). NIOSH uses derivatization with 2-hydroxymethyl-piperazine followed by gas chromatographic analysis. The OSHA method collects glutaraldehyde as the derivative with 2,4- dinitrophenylhydrazine (DNPH) with analysis by high-performance liquid chromato- graphy (HPLC). The OSHA method was selected for further evaluation as it appeared to offer a better detection limit than the NIOSH technique (4.4 ppb rather than 165 ppb for sample volumes of 15 and 4 l., respectively). The results of that evaluation are reported in this study.

Chemicals

MATERIALS AND METHODS

The chemicals used were glutaraldehyde (50% aqueous Aldrich Chemical Co; 25% aqueous, grade 1, Sigma Chemical Co.), 2,4 DNPH (AR, Aldrich Chemical Co.),

223

224 J. Cuthbert and J. Groves

hydrochloric acid (AR, FSA), phosphoric acid (HiPerSolv, BDH), acetonitrile (HPLC grade, Rathburns), ethanol (AR, James Burroughs), ammonia solution (AR, FSA), water (distilled and demineralized or reverse osmosis purified).

HPLC equipment and conditions HPLC analysis was carried out with a Waters 6000A solvent delivery system, a

Rheodyne 7125 injection valve and a Waters lambda max model 481 LC spectrophotometer at 360 nm. The column was a 25 cm Zorbax CN 5 pm, 4.6 mm i.d. The mobile phase was 55% acetonitrile, 45% buffer pumped at 2.0 ml min-‘. The buffer consisted of 1% sodium dihydrogen phosphate dihydrate adjusted to pH 5.5 with sodium hydroxide. The eluent used in the OSHA method consisted of acetonitrile, water and phosphoric acid (55%:44.9%:0.1%) but it was observed during this study that the glutaraldehyde-DNPH derivative was unstable in this solution and hydrolysed back to the starting materials with a half-life of about 90 min. No such instability was observed with the replacement eluent, which has the added advantage that it also resolves the two isomers of the glutaraldehyde-DNPH derivative which had previously eluted together (Fig. 2).

Calibration standards Glutaraldehyde is available only as aqueous solutions supplied in various

concentrations usually of nominal concentration 25 or 50% glutaraldehyde. The published OSHA method recommends a titration to determine the glutaraldehyde concentration in the stock solutions used for calibration purposes but the method was found in this laboratory to give poorly defined end-points. The following alternative titration gave a much sharper endpoint.

Potassium hydroxide (16.4 g) was dissolved in 500 ml of water; 150 ml of this solution was added to a solution of 10 g of hydroxylamine hydrochloride in 200 ml of water. One gramme of 50% glutaraldehyde solution was diluted to 100 ml with water and 20 ml aliquots added to 25 ml of the aqueous hydroxylamine solution. These were allowed to stand in the dark for 1 h and then back-titrated against 0.1 M hydrochloric acid using screened methyl orange as the indicator. The mean difference between the blank and sample titre was 20.1 ml equivalent to 49.9% glutaraldehyde.

An alternative to titration is to calibrate with the glutaraldehyde-DNPH derivative itself. This was the technique used throughout this study. The glutaraldehyde-DNPH derivative was prepared as follows. DNPH (6 g) was dissolved by heating in 100 ml of 4 M hydrochloric acid. This solution was filtered then left to cool. Pale yellow needle- shaped crystals of DNPH . HCl formed. These were washed with acetonitrile and then with diethyl ether and dried. This material in turn was recrystallized from 70 ml of 4 M HCl, and then washed and dried as before, producing about 3.5 g of purified material. Two grammes of this were added to i0 ml of concentrated HCl and diluted with 180 ml of ethanol. The mixture was warmed and stirred to dissolve the reagent. 2 ml of a 25% solution of glutaraldehyde (containing approximately 0.5 g of glutaraldehyde) mixed with 10 ml of ethanol were then added to the solution. The derivative formed rapidly and, after cooling, was filtered off. It was purified by recrystallization from boiling acetonitrile, washed with ether and dried. Analysis of the crystalline derivative gave carbon 44.44%, hydrogen 3.71% and nitrogen 24.29% against the theoretical 44.35% carbon, 3.50% hydrogen and 24.34% nitrogen. The derivative had a melting point of

The measurement of airborne glutaraldehyde by high-performance liquid chromatography 22s

Peak Area

0 0.4 0.8 1.2 1.6 2.0 2.4

Glutaraldehyde Concentration pg/2ml

Fig. 1. Calibration line for glutaraldehyde-DNPH derivative.

184-l 85°C. An attempt was made to determine the mass spectrum of the product but none could be obtained because the material has a low vapour pressure and is unstable at high temperatures.

Real samples would contain a large of excess of DNPH and also phosphoric acid. Concern was expressed that this may affect the chromatography and thus make the calibration invalid. By comparing the HPLC detector responses for two solutions, one ofjust the derivative and the other containing DNPH and acid, this was shown to have no effect on the chromatography.

Method of calibration Six samples of the purified derivative of mass between 0.04 mg and 0.25 mg were

weighed, transferred to separate 50 ml volumetric flasks and made up to volume with acetonitrile. These solutions were used directly as calibration standards and resulted in injections into the HPLC of about 2-10 ng of glutaraldehyde assuming a 10 ~1 injection loop (4.5985 pg of glutaraldehyde-DNPH derivative is equivalent to 1 pg of glutaraldehyde). Using the sum of the area under the two isomer peaks, the calibration line shown in Fig. 1 is obtained. Since the sample is desorbed into 2 ml a concentration unit of pg per 2 ml was used to give the mass of glutaraldehyde in a sample directly. A typical chromatogram is shown in Fig. 2.

Attempts to separate the isomers by fractional crystallization resulted in mixtures of the isomers in various ratios. Analysis of these mixtures by HPLC showed them to have the same response per unit mass suggesting that the U.V. response of the isomers is identical and justifying the use of a combined calibration line for the isomers. The U.V. absorption peak of the fractions shifted about 5 nm to longer wavelengths as the

J. Cuthbert and J. Groves

1

I

0 I I I I I

2 4 6 8 10

Retention Time (minutes)

Fig. 2. Chromatogram of glutaraldehyde-DNPH derivative showing two isomers. Peak identification: 1, excess DNPH reagent; 2, cis-trans glutaraldehyde-DNPH; 3, trans~rans glutaraldehyde-DNPH.

proportion of the later eluting isomer increased. The longer wavelength of maximum absorbance plus steric hindrance considerations suggests that the later eluting isomer, which is also the one in the greater proportion, is the trans-trans isomer. The earlier eluting component is probably the cis-trans isomer. The cis-cis isomer is not observed with this solvent system. It may not be present or could possibly elute with one of the other two isomers.

Generation of test atmospheres of glutaraldehyde Test atmospheres were prepared by two methods. In the first method, dry air was

passed at 1 1. min- ’ through a series of three Drechsel bottles. The first bottle contained water, the second contained M/10 aqueous sodium chloride and the third bottle contained 2% aqueous glutaraldehyde. The first bottle saturated the air with water vapour. The salt solution in the second bottle has theoretically the same partial pressure of water vapour above it as the 2% glutaraldehyde solution in the third bottle. This ensures that the relative rate of loss of glutaraldehyde and water from the third bottle containing glutaraldehyde solution remains more or less constant and so maintains an essentially constant concentration ofglutaraldehyde in the air stream. All three bottles were maintained at 15°C to prevent condensation further along the gas lines. Humidified air at a controlled temperature was mixed with the air containing glutaraldehyde then passed to a sampling chamber where up to six samples could be collected simultaneously. With this system, concentrations of glutaraldehyde remained

The measurement of airborne glutaraldehyde by high-performance liquid chromatography 27

constant over periods of several days and a constant concentration of glutaraldehyde could be delivered to the sampling devices.

In the system described above, the actual concentration of glutaraldehyde generated did not match the theoretical concentration exactly. In the determination of recovery efficiency, a system was required which deposited a known mass of glutaraldehyde on to the samplers. This was achieved by placing a known amount of glutaraldehyde in acetonitrile solution in a glass tube attached directly to the sampling device. Air was drawn over the solution and onto the sampler until the solution had evaporated. In this way, a known mass of glutaraldehyde could be transferred to the filter. This technique has the disadvantage that while the total dose or mass of glutaraldehyde delivered to the filter was known, the instantaneous concentration of glutaraldehyde was unknown and could be quite high.

Sumpliny medium Glutaraldehyde vapour is collected in the OSHA method by drawing air through a

filter impregnated with DNPH. The glutaraldehyde vapour reacts with DNPH on the filters to form the glutaraldehyde-DNPH derivative. The filters were prepared for sampling by coating them with phosphoric acid and purified DNPH. DNPH was purified as follows.

Five grammes of DNPH were converted to the hydrochloride by dissolving in 100 ml of hot 4 M HCl. The hot liquid was filtered through a sintered glass funnel, cooled and the hydrochloride crystallized as pale yellow needles. After recrystallization from 100 ml of 4 M HCl, the pure salt was filtered off and powdered, then added to a solution of 10 ml of 25% w/v ammonia in 50 ml of water with stirring to convert the hydrochloride to the free base. The DNPH was filtered off, washed with water and recrystallized from boiling acetonitrile containing a little ammonia followed by a further recrystallization from acetonitrile alone.

The filter coating solution was made by making 100 mg of purified DNPH and 0.5 ml of phosphoric acid (85%) up to 25 ml with acetonitrile. Glass fibre filters (Whatman GFA) of 37 mm dia. were supported by their edges on the rim of 30 ml beakers and 0.5 ml of filter coating solution pipetted onto them. The filters were left in a fume cupboard until the solvent evaporated and they were then ready for use. The prepared filters varied in appearance from batch to batch some being plain yellow others having flecks of red but this had no effect on their performance. Filters were stored before use in a covered container such as a Petri dish.

During sample collection, the filters were mounted in Gelman plastic air monitoring cassettes for 37 mm filters (part number 4339) or Millipore filter holders for 37 mm filters (part number MOO 037 AO).

EVALUATION OF THE METHOD

Extraction of’ the jilters The OSHA method recommends recovery of the glutaraldehyde-DNPH derivative

from the filter by extraction on a tube rotator for 1 h but gives no data to support this approach. This was assessed by sampling glutaraldehyde at approximately the OES (0.2 ppm) onto a filter, desorbing by the addition of 2 ml of acetonitrile then repeatedly injecting the solution into the HPLC at various time intervals. In between injections,

.I. Cuthhert and J. Groves

Glutaraidehyde Recovered (pg)

0.5 c

. * + +. . +

.

00 0 20 40 60 80 100 120

Time (minutes)

Fig. 3. Recovery of glutaraldehyde from three filters as a function of time.

the solution was allowed to stand on the bench. Two further filters were treated in the same way. The results are shown in Fig. 3. Up to about 40 min, there is some scatter between the three data sets although the data on the individual filters show a trend for the concentration of glutaraldehyde to decrease slowly with time from an initial high value to an equilibrium value. The filters were deliberately handled with the minimum of agitation of the solution to prevent the detachment of glass fibres from the filters which can block the HPLC injector. Some of the scatter shown in Fig. 3 in the period up to 40 min after adding acetonitrile may be due to inhomogeneous mixing. It is therefore possible that the waiting time could be shortened but if the sample is treated in the way suggested then a minimum waiting time of 40 min is recommended.

Ef3ct offlow rate and humidity on sampling eJiciency The OSHA method included a study of the effect of relative humidity on sampling

efficiency but the lowest humidity included was 33% at 30°C. This is equivalent to about 58% at 20°C. As humidity can affect the sampling efficiency onto impregnated filters, the performance of the method at humidities and temperatures more typical of those likely to be found in the U.K. needed to be assessed.

About 1.6 pg of glutaraldehyde (equivalent to sampling an atmosphere at the OES for 10 min at 200 ml min-‘) was sampled onto filters using the constant dose apparatus using air of known humidity and at three sampling rates. Three samples were collected at each combination of humidity and flow rate. The flow rates were 200,500 and 1000 ml min- ’ and the humidity values were 0, 10,20,30,50,60 and 70%. Each filter was analysed for glutaraldehyde and the results are given in Table 1.

The measurement of airborne glutaraldehyde by high-performance liquid chromatography 229

Table 1. Effect of relative humidity and sampling rate

Flow rate (ml mm’) 0 10

Overall Relative humidity mean

20 30 50 60 70 CV%*

200

Mean CV%”

500

Mean CV%

1000

Mean CV%

Mean

0.728 1.565 1.855 1.031 1.954 1.988 0.923 1.660 1.824 0.894 1.736 1.889

17.18 11.70 4.61

1.068 1.831 2.143 0.801 1.292 I.883 0.553 1.568 1.924 0.807 1.564 1.983

31.90 17.20 7.04

1.047 1.778 1.841 0.968 I .479 1.649 1.013 1.590 1.738 1.009 1.616 1.743 3.93 9.35 5.51

0.903 1.639 1.871

I.873 1.683 2.056 I.631 1.941 1.946 1.785 1.755 1.573 I .776 I .793 1.859 6.94 7.42 13.60

1.895 1.948 1.750 1.676 2.000 1.749 1.605 2.012 1.620 1.725 1.987 1.706 8.76 1.71 4.38

1.888 1.829 1.740 1.599 1.584 1.747 1.939 1.972 1.924 1.809 1.795 1.804

10.10 10.90 5.76

I.765 1.858 1.789

1.961 1.995 1.915 1.957 I.885 2.05 7.68

2.113 1.812 1.868 1.931 I .867 8.29 8.86

1.949 1.781 1.730 1.820 1.794 6.30 7.01

1.902

Results are in ng of glutaraldehyde. *Omitting results at 0 and 10% RH.

Examination of the mean recovered glutaraldehyde at each humidity in Table 1 shows that between 20 and 70% relative humidity, the capture efficiency appears to be independent of relative humidity. At 10% relative humidity, the capture efficiency shows some signs of decreasing relative to the higher humidities and at 0% relative humidity the reduction is quite distinct. The results of varying the flow rate are less clear. The final column in Table 1 giving the mean values suggests there may be a slight reduction in sampling efficiency at 1000 ml min-’ compared with 200 and 500 ml min- ’ but the effect is not large. At the dose of glutaraldehyde used in this experiment, equivalent to 10 min sampling at 200 ml min- ’ at the OES, it appears that sampling can be safely carried out in the range 2&70% relative humidity using flow rates up to 500 ml min-‘. Flow rates up to 1000 ml min-l are possible with a slight reduction in sampling efficiency. The overall coefficient of variation (CV) for all values in Table 1 except those at 0 and 10% humidity is 7.9% which includes sampling at a range of humidities and flow rates in addition to recovery and analysis stages.

Storage ofjlters after sample collection OSHA examined the effect of storage after sampling on filters which had collected

samples at 72% relative humidity and 31°C. As it was not clear whether large quantities of water vapour would affect the stability of the samples, the work was repeated at lower temperature and humidity.

Filters were loaded with glutaraldehyde immediately after preparation and then stored in a plastic bag for four different storage times and in each of three environments; a dark cupboard at room temperature, on the bench in a plastic bag and under refrigeration at about 4°C. In all cases a nominal 1.6 pg of glutaraldehyde was added to the filters from the vapour phase using air at about 20°C and 50% relative humidity. The samples were collected at 200 ml min - ’ over about 20 min. Three filters from each

230 J. Cuthbert and J. Groves

Table 2. Effects of storage at various temperatures and for various ttmcs

Mass glutaraldehyde recovered (,lg)

Storage time I day I week 2 weeks 4 WCCk>

Location* A B C A B C A B C‘ A B C 1.51 1.51 I Sl 1.55 1.60 I.66 1.47 I.57 I.58 I.67 I .46 0.86t 1.29 1.55 1.50 I.21 1.56 1.55 1.46 1.37 I.61 1.61 I57 1.52 1.62 1.60 1.67 1.37 l.16t 1.55 1.70 1.48 1.66 I.71 I .37 1.60

Mean 1.40 1.55 1.58 1.38 1.44 1.59 1.54 1.47 I.61 1.66 1.47 1.33 CV% 11.40 2.90 5.41 12.40 16.70 4.00 8.80 6.80 1.57 3.03 6.83 29.60

*Storage locations: A. stored in the dark; B, stored in plastic bags on the open bench; C, stored in the fridge (about 4°C).

TProbable outliers.

Table 3. Repeatability of the analysis

Nominal concentration ppm 0.20 0.02

Mass of glutaraldehyde recovered (pg)

I I.803 2 1.859 3 1.980 4 1.966 5 I.810 6 1.775

Mean 1.866 CV% 4.7

0.158 0.160 0.174 0.151 0.150 0.151

0.157 5.8

environment were analysed for glutaraldehyde on each of four occasions selected to give storage times of 1 day, 1 week, 2 weeks and 4 weeks. The results are given in Table 2.

The mean results do not show any trend with time or storage conditions and the recoveries after 4 weeks are (within experimental error) the same as after 1 day. Consequently, samples can be stored for at least 4 weeks at room temperature after collection. Omitting the two probable outliers, the mean of the remaining values is 1.537 pg glutaraldehyde with an overall CV% of 7.4%. This CV includes sampling, storage (under three sets of conditions and up to 4 weeks), recovery and analysis stages and is similar to that reported in the previous section.

Repeatability of the method Test atmospheres of glutaraldehyde in air at 50% relative humidity were sampled at

nominal concentrations equivalent to the OES (0.2 ppm) and l/lOth the OES (0.02 ppm). Six samples were collected at each concentration at 200 ml min- ’ over a period of 10 min. The results of analysing for glutaraldehyde are given in Table 3.

At a nominal 0.2 ppm glutaraldehyde (the current OES), six analyses gave a mean recovery of 1.866 pg equivalent to 0.223 ppm with a coefficient of variation of 4.7%. At 0.02 ppm (l/lOth the current OES) the recovery was 0.157 pg equivalent to 0.0188 ppm and the CV was 5.8%.

The measurement of airborne glutaraldehyde by high-performance liquid chromatography 231

Table 4. Calibration data for calculation of detection limit

Glutaraldehyde injected

bg) Isomer* Response (area counts) Mean CV%

0.3100 0.5037 0.74500 1.110 I.303

0.3100 0.5037 0.7450 1.1 IO 1.303

0.3 100 0.5037 0.7450 1.110 I .303

c t

c t

c-t

C-t

c t

t t

t t

I t

t-t

t 4

c-t + t--t

cct + t--t

c t+t-t

c 4 + t--t

c t+t-t

81.18 103.9 78.97 54.23 51.83 74.02 29.07 90.12 169.5 100.3 113.2 121.2 118.9 25.84

163.5 304.1 267.7 164.8 159.7 212.0 32.42 352.4 307.9 324.4 271.6 338.1 318.9 9.769 399.8 426.9 352.0 368.9 405.9 390.7 7.668

349.8 420.3 376.9 352.9 353.0 370.6 8.054 642.1 603.4 618.9 587.6 573.4 605.2 4.460 177.1 816.6 849.8 866.9 893.3 840.7 5.371

1208 1208 1236 1248 1295 1239 2.913 1416 1445 1452 1580 I444 1467 4.380

431.0 524.2 455.9 407.1 404.8 444.4 11.090 732.8 772.9 719.2 700.8 694.6 724.0 4.309 940.6 1121 1118 1032 1053 1053 7.014

1560 1516 I560 1520 1633 1558 3.037 1816 1872 1804 1949 1850 1858 3.089

*c 1. c?svrrcu~~ isomer; t&t, trans~trans isomer.

Sampler capacity A test atmosphere of glutaraldehyde in air at 50% relative humidity was sampled

with six filters at a nominal concentration of 0.4 ppm (twice the OES) for 10 min with back-up filters in place behind the main filter. At this concentration, the dilution flow in the test atmosphere was too low to provide adequate mixing in the sampling chamber so the reproducibility between samples was poor. However, the samples were analysed to provide breakthrough data. In only one case was breakthrough observed and this was less than 1%. In the remaining cases, no breakthrough was detected. The mean recovery of glutaraldehyde was 3.08 pg, equivalent to about 1.8 times the OES. The samplers should therefore have sufficient capacity to collect at least this concentration of glutaraldehyde over 10 min.

Detection limit The detection limit was determined by preparing five solutions of glutaraldehyde-

DNPH in the range 31-130 ng ml-’ and injecting each solution five times with the HPLC detector on its maximum sensitivity setting (Smith et al., 1987). The standard error of the calibration was calculated from the results for the sum of both isomers shown in Table 4 and then the detection limit calculated using the definition for detection limit of y - y, = 3S,. In this definition, y, is the detector response given by a blank solution and S, the standard deviation of the blank. The intercept of the calibration line is y, and S, is given by the standard error of the estimate of the line. Using this technique, the detection limit was found to be 0.108 ng of glutaraldehyde. A 10 ~1 injection loop was used so this corresponds to 21.60 ng in the final 2 ml used to desorb the filter. Assuming the sample is collected at 200 ml min-’ over 10 min, this equates to 0.0108 mg rne3. This is about 1/65th of the current OES. Collecting at

23’ J. Cuthbert and J. Groves

1000 ml min’ improves this to about 11300th of the OES. In each case. the quantification limit would be about five times higher than the detection limit.

FIELD TRIALS

Field trials were carried out at two hospital units using glutaraldehyde as a sterilizing agent. The trials were intended to assess the ease of use of the new method and, wherever possible, compare it with the existing method using MBTH (Turnbull, 1988).

Air samples were collected at the first hospital in a room where endoscopes were washed and sterilized after use. The endoscopes were rinsed in a sink at one side of the room and then transferred to a Keymed automatic sterilizing unit. Four pairs of samples were taken in this room. Each pair consisted of a DNPH sampler collecting at 200 ml min- ’ and an MBTH sampler operated at 1000 ml mini ‘. The sampling period was chosen to cover the cycle of cleaning, sterilizing and removing the endoscopes for re-use. Samples were also collected in a theatre preparation room while the glutaraldehyde solution used to sterilize the instruments was disposed of down a sluice. The sampling time for this operation was only 5 min. Two pairs of samples were also collected in X-ray processing areas (glutaraldehyde is used in some X-ray processing formulations).

At the second location, samples were collected in the endoscopy suite where the instruments were rinsed with water and placed in an Olympus KIO unit for sterilization. The sampling time was broadly synchronized to the turnround time of about 10 min. Most of the samples were collected in a portable sampling chamber which ensures that the atmosphere collected by the samplers is as homogeneous as possible (Ellwood et al., 1988). This allowed the simultaneous collection of two DNPH and two MBTH samples. The results thus gave a measure of both the agreement between the methods and the repeatability of each method. Four runs of this type were done, each corresponding to a cycle of the use of the sterilizing machine. Paired samples (without the use of the sampling chamber) of MBTH and DNPH were also collected in an adjoining ante room which contained a sluice for disposal of the spent solution and rinsing water. The results of the analyses are given in Table 5.

The ‘dry’ sampler used in the DNPH method proved, as expected, to be much more convenient to use than the liquid containing impinger required for the MBTH method.

The MBTH method at hospital 1 appears to have measured an essentially unvarying background response equivalent to about 0.06 mg m -3 of glutaraldehyde. The equivalent results using the DNPH method were generally at concentrations about half or less than those of the MBTH method. There does not appear to be any correlation between the two sets of results.

At hospital 2, the pairs of samples collected in the sampling chamber show reasonably good agreement between duplicate samples of the same method whereas the paired samples taken in the ante room without the sampling chamber show much poorer agreement between samples of the same type. This demonstrates the usefulness of the sampling chamber to ensure all samples are exposed to the same atmosphere.

The correlation between the two methods, however, is again poor. Apart from the first group of samples taken above the sterilizer, the DNPH results tend to be half or less than the corresponding MBTH results.

The measurement of airborne glutaraldehyde by high-performance liquid chromatography 233

Table 5. Airborne giutaraldehyde concentrations at two hospital sites

Glutaraldehyde (mg m-j) MBTH DNPH Hospital Locations

0.06 0.036 0.07 0.028 0.06 0.065 0.06 0.027 0.06 0.010 0.10 0.304 0.05 N/D 0.08 0.031

0.04 0.035 0.29 0.069

0.13 0.214 0.13 0.184

0.09 0.034 0.10 0.052

0.12 0.009 <O.Ol 0.016

0.07 0.031 0.07 0.021

1 I 1 1 1 I 1

2 2

2 2

2 2

2 2

2 2

Above sluice in endoscopy suite End of bench opposite disinfector On top of disinfector Bench opposite disinfector Theatre 2 preparation room, above sluice Theatre 2 preparation room, adjacent to sluice Beside X-ray processing equipment Beside accident and emergency department X-ray

Behind disinfector in ante room Behind disinfector in ante room

Shelf above disinfector*

Shelf above disinfector*

Shelf above disinfector*

Shelf above disinfector*

*Four samples collected with sampling chamber.

CONCLUSIONS

The DNPH method provides a method for measuring glutaraldehyde which is more sensitive than the calorimetric method it replaces and less likely to suffer from interferences. Laboratory evaluation showed that samples can be collected at flow rates up to 500 ml min - ’ and in the relative humidity range 20-70% without problem. Comparison with the MBTH calorimetric method during field trials resulted in a poor correlation between the methods but this is most probably due to the poor sensitivity of the calorimetric method and its susceptibility to interference. A document describing the HPLC method in detail is available from the authors.

REFERENCES

Ellwood, P. A., Groves, J. A. and Pengelly, M. I. (1988) A portable sampling chamber for field evaluation of diffusive samplers. HSE Internal Report IR/L/SP/88/03.

NIOSH (1989) Manual ofAnalytical Methods (3rd Edn). Method No. 2531, National Institute ofSafety and Health, Cincinnati, Ohio.

OSHA (1985) Analytical Methods Manual, Method No. 64. OSHA Analytical Laboratory, Occupational Safety and Health Administration, Salt Lake City, Utah.

Smith, K. A., Wood, S. and Crous, M. (1987) Rapid high performance liquid chromatographic method for the determination of propranolol in plasma. Analj,st 112, 407409.

Turnbull, G. B. (1988) Spectrophotometric determination of glutaraldehyde in the atmosphere using 3- methylbenzothiazol-2-one hydrazone hydrochloride (MBTH). HSE Internal Document.