What control organisms should I use in my anaerobic workstation?
CONTROL ORGANISMS FOR ANAEROBIC WORK STATIONS

There are many different views on the choice of control organisms for use in anaerobic workstations and consequently there is no standard approach.

Traditionally there has been a tendency to use a Pseudomonas sp as a negative control, in which case growth of the organism is considered to indicate incomplete anaerobiosis and a malfunctioning cabinet.   We do not recommend this approach primarily because growth of Pseudomonas sp gives no information about whether anaerobes are able to grow.   It is therefore more appropriate to show that the anaerobic workstation will support growth of those organisms you are attempting to isolate.

For this reason we would advocate that one of the NCCLS control strains are used namely

Bacteroides fragilis                               ATCC 25285
Bacteroides thetaiotaomicron             ATCC 29741
Clostridium perfringens                       ATCC 13124

The control strain can then be used to inoculate a plate and a metronidazole disc added to the control plate.

Growth of the control organism will clearly show the ability of the test system to support anaerobic growth.   Additionally the size of inhibition zone around the metronidazole can be measured to provide a quantitative control measure.   As metronidazole is only active under anaerobic conditions the user will therefore be alerted to possible cabinet failure should the inhibition zone size fall below pre-determined limits.

NB    It should be noted that poor growth of the control strain may be attributable to factors other than inadequate anaerobiosis, i.e. media, integrity of the inoculum.


How does the anaerobic workstation achieve anaerobic conditions?
ANAEROBIOSIS

Maintenance of anaerobiosis in anaerobic workstations is normally achieved through the reduction of oxygen by hydrogen in the presence of a palladium catalyst. The combination of hydrogen and oxygen occurs in the ratio of 2:1 by volume to form water vapour. The reaction, in the presence of a palladium catalyst, commences at room temperature. The reaction is exothermic and the heat generated amounts to a temperature rise of approximately 10°C for each 0.1% oxygen removed. The resultant water vapour is removed by the atmospheric conditioning system in the Don Whitley Scientific range of anaerobic workstations.

The efficiency of the catalyst is impaired in the presence of volatile organic compounds (volatile fatty acids) and hydrogen sulphide and when it gets wet. Anotox is present in Don Whitley Scientific anaerobic workstations and serves to remove volatiles including hydrogen sulphide consequently protecting the catalyst. For further information on Anotox see Brazier J S (1982) Appraisal of Anotox, a new anaerobic atmosphere detoxifying agent for use in anaerobic cabinets, Journal of Clinical Pathology 35, 233-238 and Pridmore A, Silley P (1995) An investigation into the performance of anaerobic atmosphere generation and detoxification methods. IX International Symposium of the Society for Anaerobic Microbiology. Cambridge UK.

To maintain stringent anaerobic conditions it is only necessary to ensure the catalyst is active and the ratio of hydrogen to oxygen is at least 2:1. Under such conditions the oxygen concentrations will be less than 5 ppm.

It must be emphasised that anaerobiosis is determined by

a) the ratio of hydrogen to oxygen
and
b) the presence of an active catalyst

If these two parameters are optimal then the workstation will produce stringent anaerobic conditions.

Data generated in situ has demonstrated that levels of <5 ppm oxygen can be achieved (in-house data). In an independent DHSS evaluation of a Whitley Anaerobic Workstation in 1986 levels of 23 ppm were recorded.

How do I disinfect the bubble trap on my anaerobic workstation(MKIII cabinets and older)?
APPROPRIATE DISINFECTANTS FOR USE IN THE BUBBLE TRAP OF WHITLEY MARK 3 AND COMPACT ANAEROBIC WORKSTATIONS

It is imperative that the choice of disinfectant for use in the bubble trap of Whitley Mark 3 and Compact anaerobic workstations is compatible with the acrylic composition of the workstation.

We recommend one of the following two disinfectants which have been used in long term testing within our own laboratory and which have been shown to have no corrosive properties.

1.    Aquasan Tablets (Bronopol base)
One tablet to be dissolved in 5 l of sterile water and added to the bubble trap and filled to the desired level.

The UK distributors of Aquasan Tablets are
Guest Medical, Enterprise Way, Edenbridge, Kent, TN8 6EW, England
Tel: +44 (0)1732 867466        Fax: +44 (0)1232 867476

They are prepared to deal directly with users of Whitley anaerobic workstations.

2.    Sodium metabisulphite
0.5 g to be dissolved in 1000 ml of sterile water and added to the bubble trap to the desired level.

The bubble trap should be cleaned before disinfectant is added.   During use the bubble trap should be checked each week and the disinfectant level topped up where necessary.


What gas mixtures can I achieve in my Whitley VA Workstation?
WHITLEY VA WORKSTATION

The Whitley VA workstation has been developed primarily for the study and isolation of microaerophilic organisms including Campylobacter spp, Helicobacter pylori and other similarly fastidious organisms.   Nitrogen, carbon dioxide, air and a 10% hydrogen/90% nitrogen mix can be combined to provide a specific atmosphere.

The selected atmosphere can be in the range:

Oxygen                       0 - 20%
Carbon Dioxide        0 - 20%
Hydrogen                   0 - 5%
Nitrogen                     Balance

The Whitley VA workstation incorporates safeguards to prevent an operator selecting a combustible combination of gases.  Oxygen and hydrogen are combustible under certain conditions so the maximum concentration of hydrogen cannot be selected to exceed 4% when the oxygen concentration has been selected to be equal to or greater than 6%.

What are the regulatory standards that the Whitley Workstation range complies with?

WHITLEY WORKSTATION COMPLIANCE

Don Whitley Scientific regularly receive requests concerning compliance of respective products currently sold by the Company. All new products are designed to meet the stringent EC regulations and consequently all carry the CE mark. This technical note specifically relates to Whitley Workstation compliance.

1. The Whitley Workstation range has been designed, developed, tested and constructed to comply with the requirements of the CE marking standards required in Europe.

2. CE marking offers manufacturers the option of self-certification where professional competence in a specific industrial sector can be demonstrated. Don Whitley Scientific has been developing anaerobic and microaerobic workstations since 1980; has supplied well over 1000 systems worldwide, and is a recognised authority in this field.

3. Where self-certification is chosen as the preferred method of compliance with the requirements of the standard, a manufacturer must assemble a comprehensive Technical Construction File (TCF). This document details such areas as the technical justification for the selection of specific components; the results of testing equipment to the requirements of specific internationally-accepted standards in areas such as electrical safety, electromagnetic compatibility considerations, etc; risk assessment under single fault conditions and many other matters.

4. The Whitley Workstation range has been subject to this approach. All documentation is filed within Product Development at Don Whitley Scientific. Testing during development was performed in accordance with BS EN 61010-1 and is documented within the TCF. By the successful completion of these tests, the equipment satisfies the requirements of the Low Voltage Directive (73/23/EEC) and the Machinery Directive (89/392/EEC). To ensure all workstations manufactured subsequently meet the requirements of these directives, the tests described in Annex K of BS EN 61010-1 are performed on each workstation produced.

5. Through experience gained during almost 20 years of designing and developing workstations; by specifying components which themselves carry the CE mark - installed and used in accordance with the manufacturer’s instructions, it is considered that the Whitley Workstation range meets the requirements of the Generic Emission Standard (Residential, Commercial and Light Industrial) - EN 50081-1, and the Generic Immunity Standard (Residential, Commercial and Light Industrial) - EN 50082-1.

6. By compliance with all the directives indicated above, the equipment may carry the CE mark.

7. We are informed that many other countries have already (or are presently putting in place) mutual recognition agreements to accept design, development and test methods used elsewhere in the world.

8. As a small manufacturing company taking all the issues described above into account, Don Whitley Scientific does not have the resources to ensure our products meet all requirements of all countries. If your country has specific requirements please advise us accordingly.

REGULATORY COMPLIANCE - WHITLEY WORKSTATIONS

The Whitley Workstation range has been designed, developed, tested and constructed to comply with the requirements of the CE marking standards required in Europe.

CE marking offers manufacturers the option of self-certification where professional competence in a specific industrial sector can be demonstrated. Don Whitley Scientific has been developing anaerobic and microaerobic workstations since 1980; has supplied well over 1000 systems worldwide, and is a recognised authority in this field.

Where self-certification is chosen as the preferred method of compliance with the requirements of the standard, a manufacturer must assemble a comprehensive Technical Construction File (TCF). This document details such areas as the technical justification for the selection of specific components; the results of testing equipment to the requirements of specific internationally-accepted standards in areas such as electrical safety, electromagnetic compatibility considerations, risk assessment under single fault conditions and many other matters.

The Whitley Workstation has been the subject of this approach. All documentation is filed within Product Development at Don Whitley Scientific. Testing during development was performed in accordance with BS EN 61010-1 and is documented within the TCF. By the successful completion of these tests, the equipment satisfies the requirements of the Low Voltage Directive (73/23/EEC) and the Machinery Directive (89/392/EEC). To ensure all workstations manufactured subsequently meet the requirements of these directives, a thorough test sequence is performed on each workstation produced.

Through experience gained during almost 20 years of designing and developing workstations; by specifying components which themselves carry the CE mark - installed and used in accordance with the manufacturer’s instructions - it is considered that the Whitley Workstation range also meets the requirements of the Generic Emission Standard (Residential, Commercial and Light Industrial) - EN 50081-1 and the Generic Immunity Standard (Residential, Commercial and Light Industrial) - EN 50082-1.

By compliance with all the directives indicated above, the equipment may carry the CE mark.

How stable is the temperature in a Whitley Workstation?
TEMPERATURE STABILITY IN A WHITLEY WORKSTATION

The development brief for the Whitley Workstation product development team required that temperature variation within the working area of the workstation would be maintained within a temperature band of ± 0.5°C.

Whilst the temperature profile across the workstation has been determined to be within  ± 0.5°C, the operating temperature is controlled by measuring the temperature of the return path atmosphere before the heater.   This ensures that the temperature measuring probe is positioned in an area inaccessible to the user and therefore not prone to accidental damage.

DWS In-House Test Procedure

1.    The Whitley Workstation is powered up and the temperature controller is set to 36.5°C.   The instrument is allowed to stabilise for 4 hours.

2.    A NAMAS-traceable, calibrated mercury/glass thermometer placed in a bottle of glycerol is positioned centrally on the top shelf of the workstation, 50 mm from the front edge.   The workstation is left to stabilise for at least a further 4 hours.  The thermometer reading is noted.

If necessary the display on the MACS temperature controller is adjusted during the manufacturing process to match the thermometer reading.

Should there be a difference between the controller and actual workstation temperature readings, follow this procedure to introduce the required offset:

Press and hold ? and ‚ for 3 seconds.
Press ‚ once to Level 1.
Press and hold × and press ? twice to Level 3
Press ? seven times until Zero is displayed     

Press and hold × and press ‚ or ? until the required difference in temperature is displayed eg -2.0 or 2.0.

Press and hold ‚ and ? for 3 seconds to return to the temperature display.

The temperature display will now decrease or increase by the value which has been entered.   The controller and thermometer readings within the workstation will now correlate.

3.    Product Development should be informed if the difference between the set point of the temperature controller and the thermometer is more than 1°C.

How does the Whitley Workstation and airlock mix the correct gas mixture?
MECHANISM OF GAS MIXING

To help understand the possible gas mixtures within the Whitley range of anaerobic workstations the following has been compiled to explain the fundamental differences.

Whitley MG Workstation

The Whitley MG500 and MG1000 workstations have been designed to operate from a bottle of anaerobic gas mixture. This is a world-wide industry standard gas mixture consisting of 80% Nitrogen, 10% Hydrogen and 10% Carbon Dioxide. This gas mixture is connected to the workstation and introduced into the chamber on demand. A pressure switch is set to maintain a small positive internal pressure. Any pressure drop within the workstation permits gas to flow into the unit. In addition, a forced gas injection timer delivers a small volume of gas at pre-determined intervals. This ensures that fresh gas is always being delivered into the chamber thus helping to maintain anaerobic conditions.

For applications where knowledge of gas composition is critical, some gas bottle suppliers can provide certification of the contents of an individual gas cylinder - although a premium price may apply.

Whitley MG Workstation + TG Airlock

The TG Airlock has been designed as part of the modular Whitley Workstation system. It can be bolted onto any MG workstation. As with the MG workstations, gases are delivered via a pressure switch, however the flow rates and gas mixing methods are different. This configuration permits the use of three separate cylinders of Nitrogen, Hydrogen and Carbon Dioxide. Nitrogen and Carbon Dioxide are combined and delivered together. Fail safe mechanisms ensure that Hydrogen can only be delivered as a function of Nitrogen flow. If the Nitrogen flow were to fall below a pre-set safe level, Hydrogen would no longer be injected into the chamber, ensuring that a level of 10% Hydrogen can never be exceeded. This method of gas delivery can significantly reduce running costs where cylinders of pre-mixed anaerobic gas are expensive.

As with the basic instrument, a forced gas injection timer also delivers gas at pre-determined intervals. This ensures that fresh gas is always being delivered into the chamber thus helping to maintain anaerobic conditions.


What is the proportion of gasesnin premixed anaerobic gas mixture?
ANAEROBIC GAS MIXTURE

Mixed gas anaerobic workstations manufactured by Don Whitley Scientific Limited operate on a supply of anaerobic gas consisting of 10% Hydrogen, 10% Carbon Dioxide and 80% Nitrogen, (known as a 10:10:80 mixture).

Don Whitley Scientific Limited has been a manufacturer of anaerobic workstations since 1980. More than 1000 units have been supplied worldwide. The majority operate on the 10:10:80 mixture. There have not been any incidents reported regarding the use of the 10:10:80 gas mixture on any Don Whitley Scientific anaerobic workstation.

Whilst the 10:10:80 mixture is correctly classified as an extremely flammable gas, very specific conditions apply before there is an explosive potential:

1. Anaerobic gas mixture needs to represent between 44% and 76% by volume in air, thereby providing a minimum of 5% concentration of oxygen by volume to sustain combustion.

2. A source of ignition
It is however important at this point to emphasise the difference between the terms “explosive” and “flammable”. If a flammable mixture is ignited in an open space it will simply burn. However, if a flammable mixture is ignited in an enclosed space it may cause an explosion.

Strictly speaking it is therefore incorrect to describe a gas as explosive as the explosiveness depends on both the gas and its surroundings.

The few parts per million of oxygen present in a fully commissioned and operational anaerobic workstation cannot create a hazardous situation within the workstation. The transition from aerobic to anaerobic conditions (during the commissioning activity) does include a brief crossover period when an explosive mix is present inside the workstation. If the workstation is first flushed with a 100% nitrogen mixture any risk is completely eliminated.

Anaerobic workstations produced by Don Whitley Scientific must always be installed and commissioned by our own technical staff or by those who have received full training from Don Whitley Scientific. Our workstations must always be operated in accordance with the instructions contained in the User Manual.

Leakage from anaerobic gas cylinders into a confined airspace (for example a poorly ventilated small room) could represent a hazard. A 10:10:80 mixture is also an odourless asphyxiant. The provision and maintenance of the gas supply up to the point where it enters the workstation is the responsibility of the user.


Can I use Virkon in my anaerobic workstation?
THE USE OF VIRKON IN MICROBIOLOGY WORKSTATIONS

After a thorough study lasting 20 weeks we have recently concluded that the presence of any uncovered containers of Virkon in both anaerobic and variable atmosphere workstations leads to the unacceptable degradation of stainless steel, brass and some other metal components. The effect is more considerable in variable atmosphere workstations – probably due to the presence of oxygen in the selected gas mixture. These conclusions are supported by technical information available on the Virkon manufacturer’s website and independent observations by an Institute of Materials metallurgist.

In two instances we believe the presence of Virkon vapour led to the premature failure of porthole inner door springs.

If Virkon is the preferred sanitising solution, these adverse effects are minimised by ensuring that Virkon within any workstation is always kept in a covered vessel when not in use.

We feel it appropriate to draw this matter to the attention of our distributors and customers.


How do I transfer items using the Whitley Workstation airlock?
AIRLOCK TRANSFER OF ITEMS INTO THE WHITLEY WORKSTATION

Most Whitley Workstations incorporating an airlock are used to transfer quantities of petri dishes to and from the main chamber. The airlock uses a transfer tray system to make sample transfer as easy as possible.

Dimensions

a. With a full length shelf in place in the workstation chamber

A tray measuring 295 mm x 295 mm, accommodating up to 90 x 90 mm Petri dishes can be transferred into the chamber. There is also still room for a small amount of loops, swabs etc.

b. Without the full length shelf in place in the workstation chamber

An object measuring up to 295 mm long x 295 mm deep x 285 mm high can be passed through the airlock.

Tall Items

The airlock maximum internal height is 350 mm, permitting flasks and other tall items to be transferred into the chamber (even though the inner door opening is only 290 mm high) by tilting the vessel as it passes through the doorway.

How do I check if my workstation is anaerobic?

WHITLEY WORKSTATION BUBBLER

In response to customer feedback DWS have developed the “Whitley Bubbler” as an accessory to the Whitley range of workstations. The bubbler simply pumps the internal atmosphere of a Whitley workstation through a methylene blue indicator solution which when clear indicates anaerobiosis. The indicator will turn blue in the presence of oxygen. Whilst an effective measure of oxygen the indicator is responding to a change in redox potential. With methylene blue at pH 7 and 37°C an Eh value of -28 mV corresponds to a 95% reduction of the blue oxidized form; the absence of any apparent colour should ensure that a value below this has been attained, and in normal circumstances this indicates successful removal of oxygen.

The pump is activated from outside the workstation and so readings can be taken at steady state and are not influenced by oxygen taken into the workstation upon entry.

Indicator Solution

*  The dry 2 part indicator solution mixture has a one year shelf life.

*  Distilled water should be added to the component mixes as indicated on the label. The separate hydrated solutions have a shelf life of six months at 4°C.

*  The mixed indicator solution should be changed at least monthly, or if it turns brown.

*  Never add new solution to old, discard the old solution, prepare the new solution and reduce by boiling.

*  Anaerobic Indicator solution has a DWS Part Number A00038.

What are the growth conditions necessary for Helicobacter pylori in a Whitley Workstation?

Optimal atmospheric conditions for growth of Helicobacter pylori have not been clearly defined. In a DWS study presented at the 9th International Workshop on Campylobacter, Helicobacter and Related Organisms held in 1997 in South Africa we showed that the addition of 3% hydrogen to the incubation atmosphere greatly enhanced growth of Helicobacter pylori as assessed by colony size.

 

Henriksen et al (2000) have published a more detailed study of the assessment of optimal atmospheric conditions for growth of Helicobacter pylori in the European Journal of Clinical Microbiology.

 

In this study the number and diameter of colonies of Helicobacter pylori isolates growing on agar plates were determined to compare five methods that produce a culture atmosphere. No catalyst was applied. No significant difference was found between two hydrogen-based kits that have a different capacity for production of H2. These hydrogen-based methods were significantly better than all others evaluated, including one kit that produces ascorbic acid that binds with oxygen. Growth was significantly improved when the atmosphere outside the plastic incubation jars was enriched with 10% CO2, but carbon dioxide enrichment alone (ie no reduction of the oxygen concentration) gave a very poor yield. The colony diameter was a sensitive and reliable measure of atmospheric conditions, as the mean intra- and interobserver difference between repeated readings was £ 0.1 mm for 82% and £ 0.2 mm for 95% of the isolates.

 

This study thus supported the initial DWS findings. An additional observation was that in all cases there was leakage of gas across the jars.

 

It is therefore clear that the use of a Whitley VA Workstation with the ability to alter the respective CO2, O2 and H2 concentrations provides significant advantages to laboratories working with Helicobacter pylori. Variable atmosphere workstations are used within the UK Helicobacter Reference Laboratory based at the Colindale Public Health Laboratory.

 

Reference

Henriksen T-H, Lia A, Schøyen R, Thoresen T, Berstad A. Assessment of Optimal Atmospheric Conditions for Growth of Helicobacter pylori. European Journal of Clinical Microbiology and Infectious Disease (2000) 19: 718-720

How do I clean my Whitley Workstation?
DECONTAMINATION AND CLEANING WHITLEY WORKSTATIONS

DWS recommend that as part of the daily checks users ensure that the workstation is free from spillage. Cleaning should be carried out as necessary. Prior to servicing it may be necessary for the chamber to be decontaminated.

This Technical Note describes the procedure for cleaning and decontamination and details the course of action should any spillage not be contained in the chamber base tray.

CLEANING AND DECONTAMINATION

The transparent and/or white acrylic on the inside and outside of the Whitley  Workstation range may be swabbed with a 1% solution of Labdet 100 (DWS) in warm water and dried afterwards with a soft clean cloth. In the case of culture spillages then a 5% Hypochlorite solution should be applied to the spillage and left for 30 minutes. It should then be mopped up and the surface swabbed with 1% Sodium Thiosulphate solution. As an alternative "VIRKON" can be used as a decontaminant at a 1% (w/v) concentration. VIRKON should not be left inside the workstation in an open container - for more information see Technical Note MA25.

If hypochlorite solution is being used inside the chamber it is advisable to remove the "Anotox" and Deoxo 'D' Catalyst sachets.

Never use any solvent on the acrylic surfaces of the workstation. Use only water and a mild detergent solution (ie Labdet 100 1% solution) as a cleaning agent.

If spillage is not contained in the base tray then access underneath the tray is achieved by following the steps below:

1. The hinged floor is raised by unfastening the studs on either end panel (see figure 6.10). On an MG1000 workstation there is also a retaining block on the central rib. Now lift the floor panel(s), partially closing the inner porthole doors in order to obtain the required clearance and then opening the inner doors once the floor panel(s) are rotated past them, so as to provide a rest.

2. The Anotox sachets and catalyst should be removed from their relevant positions and replaced with new sachets.

Scratches on the acrylic plastic may be removed by gently polishing the surface with 'DURAGLIT WADDING' followed by wiping with a soft clean cloth.

Deep scratches may require the use of Wet and Dry abrasive paper used wet, followed by polishing with 'DURAGLIT' - seek advice from Don Whitley Scientific Limited or our authorised agents overseas.

How is the gas flow fault on a Whitley Workstation initiated?
The Whitley range of anaerobic workstations operate at a slight positive pressure thereby ensuring the integrity of the selected atmosphere inside the chamber. If there were to be a leak in the chamber then there would be a pressure drop which in turn would result in a call for gas. If there is a continuous flow of gas for five minutes then a Flow Fault error will be initiated. Under these conditions the green gas demand LED on the front control panel will be lit. Under normal conditions this LED should illuminate briefly every few minutes.

ACTION

If the flow fault is showing the following checks should be made:

1. Check that both inner porthole doors are securely closed.

2. Check that the oil in the oil bottle is at the correct level.

If the above checks are OK it may be that there are a number of small leaks around the end plate screws, which amounts to a large leak. If this is the case these can be tightened using a 10 mm spanner. TAKE CARE NOT TO OVER TIGHTEN THESE PLASTIC NUTS. If the leak is still apparent, tie a knot in both sleeves and watch to see if they inflate. If they do, there is a leak around the portholes, which will probably require an engineer to visit.

How does the Whitley Workstation wireless footswitch work?

OPERATION OF WHITLEY WORKSTATION FOOTSWITCH TRANSMITTERS

The porthole door system on the Whitley range of workstations operates through an advanced wireless control technology.

In specifying a radio control system suitable for use in hospitals and similar establishments it was important for us to choose a system not susceptible to external interference. Our system uses similar principles to those in car and domestic security alarms. If the receiver detects a transmitted signal that it does not recognise (even, for example, transmitted from another footswitch operating on a different channel) at the same time as a legitimate signal is detected, it will not perform the task as instructed. This is a safety feature. The implication is that two or more workstations installed near one another cannot have the porthole sequences performed at exactly the same time as each other. When we explain to customers the thinking and reasoning behind this safety feature they see the wisdom of how the system works. If a customer does not wish to use the radio control system then a hard wire option for operating the footswitches is available.

The Whitley Workstation range is designed to permit more than one workstation to be placed near each other. When sales are made to distributors we tend not to know who the end customer is or where the workstation will be installed so we always set the same default recognition code on every transmitter and receiver. The default code consists of switches 1-4 “on” and switches 5-8 “off”.

QUESTION

We have a customer who has two workstations in the same laboratory and the footswitch for one is controlling the function on the second cabinet. Can I change the footswitch settings?

ANSWER: YES

PROCEDURE
1. Remove the top cover from the foot-switch (4 screws).
2. Locate the radio transmitter (small PCB) and remove the plastic screw. Expose the top of the PCB and locate the row of 8 dip switches. These will be factory set to 1-4 on, 5-8 off. Adjust 1 of the switches and take a note of the new setting. Photo 2.
3. Reassemble the foot-switch but do not over tighten the plastic screw.
4. Remove the lid from the workstation top box and locate the radio receiver (front right hand corner).
5. Set the radio receiver code switches to the same setting as the transmitter.
6. Check that the unit is functioning by testing the foot-switch.

How do I filter the exhaust gases from a Whitley Workstation?
WHITLEY WORKSTATION - FILTRATION OF EXHAUST GASES

There are instances when customers wish to filter the exhaust gases from a workstation. This would normally only occur after a laboratory has carried out a Risk Assessment and identified a need to prevent the potential egress of hazardous organisms into the laboratory environment.

MODIFICATIONS

The Whitley range of workstations can be modified to provide for filtration of the exhaust gases. In principle the output gases from the pump and the oil equilibration system are filtered. Additionally the evaporator is replaced by a separate sealed condensate system which has to be emptied manually.

Workstations with this modification operate on non-standard software such that an airlock exit cycle has to be run before plates can be removed from the airlock. This flushes the airlock with nitrogen thereby diluting the microbial load within the airlock.

It is important that the customer also considers the laboratory protocol to be used in conjunction with operating the sleeves as they will need to be fully flushed with nitrogen each time the cabinet is exited.

Is it possible to run a Whitley Workstation at lower temperatures?
The normal operating temperature of the Whtitley Workstation can be set between 5°C above ambient temperature and 42 ± 2°C.

The standard refrigeration unit is designed to operate at 8 ± 1.5°C when located in an ambient temperature of 21 ± 2°C. Other set temperatures are achievable, but because accurate temperature control is influenced by so many interdependent factors, discussion must take place with our product development staff if a specific installation is required to operate within conditions different from those stated.

There are some general principles to be aware of when considering refrigeration options.

TECHNICAL CONSIDERATIONS

1. Does the user require the workstation to operate at a fixed temperature or is it likely that they will want to regularly change the operating temperature?

2. Is the receiving laboratory air conditioned? If the laboratory has a variable air temperature according to the time of day or season it will not be possible to control and accurately maintain the internal temperature of the workstation across a fluctuating ambient temperature range. DWS staff need to be informed of the temperature variability within the laboratory prior to accepting an order.

3. Low temperature operation may also generate moisture on the outer surfaces of the workstation as this is likely to be one of the coolest surfaces in the laboratory and will act as a condenser panel for atmospheric moisture.

How do I know if my workstation is leaking?
GAS DEMAND IN WHITLEY WORKSTATIONS

Customers who have received a Whitley Workstation and have had no previous experience of anaerobic workstations can sometimes feel concerned about the rate of gas usage within the workstation.

As a guide to gas usage it can be useful to observe the green gas demand LED on the workstation front control panel. When there is nobody working in the chamber the green gas demand LED should come on no more than every 5 to 10 minutes. Under some circumstances it may even be 15 minutes, this is acceptable.

If the gas demand is more frequent than every 5 minutes it is advisable to check for gas leaks and discuss the issue with your DWS engineer/distributor.

How many plates can I get in my Whitley Workstation?

INCUBATION CAPACITY OF WHITLEY WORKSTATIONS

Don Whitley Scientific quotes incubation capacity based upon the number of 90 mm petri dishes that can be accommodated within the workstation, whilst still providing a generous working area. The figures should be viewed as the maximum capacity available.

The figures should be reduced by 90 x 90 mm Petri dishes if an airlock is fitted. This reduction in capacity provides the space necessary for the airlock tray to be pulled into the chamber.

MG500 540 x 90 mm Petri dishes
VA500 540 x 90 mm Petri dishes
MG1000 1080 x 90 Petri petri dishes


Internal Dimensions of the MG500 and VA500

Incubation Area: Length: 840 mm Depth: 330 mm Height: 440 mm

A removable shelf 10 mm thick divides this space into two areas, the lower of which is 220 mm high, the upper of which is 210 mm high.

Working Area: The working area is 840 mm wide ´ 160 mm deep with a height of 330 mm at the front of the chamber, rising to 440 mm immediately before the incubation area.

Internal Dimensions of the MG1000

Incubation Area: Length: 1680 mm Depth: 330 mm Height: 440 mm

A removable shelf 10 mm thick divides this space into two areas, the lower of which is 220 mm high, the upper of which is 210 mm high.

Working Area: The working area is 1680 mm wide ´ 160 mm deep with a height of 330 mm at the front of the chamber, rising to 440 mm immediately before the incubation area.


NOTE: Please check with Don Whitley Scientific if equipment to be housed inside a workstation appears to exceed the stated dimensions as a tolerance has been applied to the dimensions. Don Whitley Scientific will work with distributors or customers to modify workstations to meet specific needs.

How does the palladium catalyst work?
The DWS range of anaerobic workstations utilises a palladium catalyst for removal of oxygen from the incubation atmosphere. In the presence of palladium catalyst, residual oxygen is reduced by hydrogen with the production of water. The water is automatically removed from the workstation.

Under normal in-use conditions the catalyst will remain effective for up to 6 months. At 6 months it is recommended that the catalyst sachet is replaced by a new sachet . It is the practice of some laboratories to “recharge” the catalyst by heating at temperatures of 160°C. This will indeed improve the catalyst life by driving off moisture and freeing up active sites on the catalyst but it will not fully recharge the catalyst. DWS cannot guarantee maintenance of anaerobiosis within a workstation after 6 months unless new catalyst sachets are being used.

Catalyst sachets will remain active and viable for a minimum of 2 years when kept dry within the supplied polythene bag.

Product codes for catalyst sachets are dependent upon the workstation in which the sachet will be used:

Compact Workstation A01540
Mk 3 Workstation A00106
WISE Workstation A00212
Whitley Workstation A02130

How can I get single plates into a Whitley Workstation without using the ports?
WHITLEY WORKSTATION SINGLE PLATE ENTRY SYSTEM

A single plate entry system can be fitted to Whitley Workstations to enable rapid transfer of single petri dishes and other small items.

The dimensions of the single plate entry system are:

Width 98 mm and height 20 mm.

The single plate entry system will allow 90 mm diameter petri dishes and 98 mm square plates and standard 96 well microtitre plates to be passed into the workstation.

NB This option can only be added when the instrument is being built and therefore needs to be specified at placement of order.

DWS Product Code - A02711

How much gas does the Whitley DG250 Workstation use?
The Whitley DG250 anaerobic workstation can operate on an anaerobic gas mixture (10% CO2, 10% H2, 80% N2) and nitrogen gas or on anaerobic gas mixture alone. Projected gas usages have been calculated as shown below.

Assumptions

The following have been used to calculate gas usage

a) A normal operating gas requirement of four calls per hour and two timed gas injections

b) Three entries into the workstation per working day (assuming the sleeves are evacuated twice and a seven day working week).

c) Anaerobic gas mixture cylinder volume 9550L

d) Nitrogen gas cylinder volume 9780L.

Gas Usage

Operation of workstation using anaerobic gas mixture and nitrogen:

Expected life of anaerobic gas cylinder 33 weeks
Expected life of nitrogen gas cylinder 93 weeks

Operation of workstation using anaerobic gas mixture alone:

Expected life of anaerobic gas cylinder 24 weeks

NB These usage rates are based on calculation: in use rates will depend on local practices.


What do i do if my Whitley DG250 Workstation si frequently calling for gas?
If a Whitley DG250 Workstation is calling for gas more often than once every five minutes, carry out the following procedure:

1. Check that the silicone tube from the condenser cover is connected to the luer fitting on the condenser plate at the rear of the unit.

2. Check that the blue tubes to the portholes are connected correctly. Connect a pair of sleeves (one to each porthole) and tie a knot in each sleeve. If the sleeves inflate this would indicate that the inner door seals are the cause of a leak. Check that the clamping mechanism is applying a slight pressure on the seals. TAKE CARE NOT TO OVERTIGHTEN.

3. Check that the large blue seal is free from obstructions, that the four shell clamps are secured and that the front and rear panels are in the correct position.

4. Check that the pipework inside the equipment enclosure is all connected.

5. Ensure the blow-off valve (a safety device to protect the chamber in the event of any over pressurisation) is intact, complete and properly assembled.

NOTE: this component is located within the equipment enclosure on the right hand side when viewed from the front.

6. Check for leaks around the letterbox/blanking plate area.

NOTE: If a gas leak detector is used to assist these checks please note that the detector is only sensitive to gas mixtures containing hydrogen.

Can I use a UV light to sterilise my Whitley workstation?
From time to time we are asked whether UV lights can be used to sterilise an anaerobic workstation. This technical note explains why this cannot be carried out as a routine procedure.

Use of UV Light

The use of a UV lamp for sterilisation purposes in any workstation which is made out of acrylic will damage the acrylic shell and therefore should not be used.

Normal cleaning procedures are described in Technical Notes MA25 and MA32 and are sufficient for routine use. If there is a need for sterilisation then formaldehyde can be used as the sterilising agent under controlled condition.

A UV “woods lamp” may be used for diagnostic purposes for determining fluorescence of certain anaerobes. This is available from DWS as code A00041, but this is not for sterilisation purposes.

It should also be pointed out that UV light will also cause damage to growing cultures of bacteria.


How do I clean my Whitley Workstation?
DISINFECTION AND CLEANING - Whitley DG250 Workstation

DWS recommends that as part of the daily checks, users ensure that the workstation is free from spillage. Cleaning should be carried out as necessary.

This Technical Note describes the procedure for cleaning and decontamination, and details the course of action should any spillage not be contained in the chamber base tray.

Cleaning and Decontamination

The transparent and/or white acrylic on the inside and outside of the Whitley DG250 Workstation may be swabbed with a 1% solution of Labdet 100 (available from DWS Stock Code D003) in warm water and dried afterwards with a soft, clean cloth. In the case of culture spillages, a 5% hypochlorite solution should be applied to the spillage and left for 30 minutes. It should then be wiped up and the surface swabbed with a 1% sodium thiosulphate solution. As an alternative, Virkon can be used as a decontaminant at 1% (w/v) concentration. Virkon should not be left inside the workstation in an open container - for more information see Technical Note MA25.

If a hypochlorite solution is being used inside the chamber, it is advisable to remove the Anotox and Deoxo ‘D’ catalyst sachets.

Never use any solvent on the acrylic surfaces of the workstation. Use only water and a mild detergent solution (ie Labdet 100 1% solution) as a cleaning agent.

Scratches on the acrylic plastic may be removed by gently rubbing the surface with Duraglit wadding and then polishing with a soft, clean cloth.

Deep scratches may require the use of wet and dry abrasive paper, used wet, followed by polishing with Duraglit - seek advice from Don Whitley Scientific Limited or their authorised agents overseas.


What do I need to know if my workstation goes aerobic?
QUESTIONS TO ASK IF AN ANAEROBIC WORKSTATION IS GOING AEROBIC

1.    How do you know your chamber is going aerobic?

    Do you use control sample culture plates?   If so, what organisms?
    Do you use anaerobic indicator strips?
    Do you use an anaerobic indicator solution?

2.    When was the catalyst last changed?
Did you change the Anotox at the same time as the catalyst?

3.    How many plates are you culturing on a daily basis?

4.    What organisms are you growing – do any of these cultures produce volatiles that could “poison” the catalyst - eg hydrogen sulphide?

5.    Are you evacuating and gassing the sleeves at least twice every time an operator enters a commissioned workstation?

6.    Is the unit being operated using 3 separate gases i.e. Nitrogen, Hydrogen and Carbon Dioxide or one bottle of anaerobic gas.   If one bottle of gas is being used does it contain 5% or 10% Hydrogen.

7.    How often does the Gas Demand LED illuminate?

8.    Do you have plastic sleeves or latex sleeves?

 9.    Are the sleeves split anywhere?

10.    Does the workstation call for gas excessively while the sleeves are being used?

11.    When was the workstation last serviced and by whom?

What is the theory behind impedance?
AOAC APPROVAL

Background

AOAC adopted as first action the automated conductance method for Salmonella in foods. The method is based on the principle of conductance changes due to the presence of Salmonella compared to those for non-salmonellae.

The Initial AOAC trial was co-ordinated by Malthus. AOAC clearly state that any other equipment that meets the critical parameters and system suitability standards of the method is acceptable, ie RABIT.

Current Practice

Principle

Samples are pre-enriched in buffered peptone water-lysine-glucose broth followed by 2-tube conductance assay in selenite-based media containing trimethylamine-N-oxide and dulcitol (Easter and Gibson) and lysine (Ogden's Lysine). Salmonella spp typically give large conductance changes in these 2 media compared to those for non-salmonellae. Presumptive positive result is obtained within 48 hours.

Confirmation of Presumptive Positive Conductance Results

Presumptive positive conductance assay indicates that Salmonella may be present. All presumptive positive results must be confirmed.

Ref: JAOAC.



Method Performance for Automated conductance method for Salmonella in Foods as published by AOAC

Food Type
Level
Method
Performance Rate

Coconut
Low

High
B
M
B
M
72.00(9.32)
72.00(7.25)
77.33(9.33)
86.67(5.04)

Fish Meal
Low

High
B
M
B
M
27.50(5.74)
36.25(7.12)
38.75(8.05)
80.75(8.56)

Prawns
Low

High
B
M
B
M
1.54(2.54)
3.07(2.08)
26.15(8.28)
23.08(7.79)

Non-Fat Dried Milk
Low

High
B
M
B
M
27.06(6.17)
29.41(6.67)
92.94(5.93)
95.29(4.72)

Liquid Egg
Low

High
B
M
B
M
87.69(4.82)
92.30(2.82)
95.38(2.43)
93.85(4.17)

Minced Beef
Low

High
B
M
B
M
100.00(0.00)
100.00(0.00)
100.00(0.00)
100.00(0.00)

An analysis of variance (ANOVA) technique applied to the data indicated that the method means were not significantly different (P > 0.05) at each level of inoculum for each food type.

B - BAM/AOAC method; M = automated conductance method.
Performance rate is shown as sensitivity (standard error in parentheses). Performance rate for specificity was 100.00 at both levels for each method for all food types listed.


What accreditation does impedance technique have?
AOAC APPROVAL

Background

AOAC adopted as first action the automated conductance method for Salmonella in foods. The method is based on the principle of conductance changes due to the presence of Salmonella compared to those for non-salmonellae.

The Initial AOAC trial was co-ordinated by Malthus. AOAC clearly state that any other equipment that meets the critical parameters and system suitability standards of the method is acceptable, ie RABIT.

Current Practice

Principle

Samples are pre-enriched in buffered peptone water-lysine-glucose broth followed by 2-tube conductance assay in selenite-based media containing trimethylamine-N-oxide and dulcitol (Easter and Gibson) and lysine (Ogden's Lysine). Salmonella spp typically give large conductance changes in these 2 media compared to those for non-salmonellae. Presumptive positive result is obtained within 48 hours.

Confirmation of Presumptive Positive Conductance Results

Presumptive positive conductance assay indicates that Salmonella may be present. All presumptive positive results must be confirmed.

Ref: JAOAC

How to I make indirect impedance cells?
PREPARATION OF INDIRECT RABIT CELLS

Background

Indirect RABIT cells are simple to prepare and allow highly sensitive detection of a wide range of microorganisms. Care in the preparation of indirect cells will ensure maximal shelf life and consistent sensitivity. The indirect impedance technique detects microbial growth on the basis of the carbon dioxide produced during normal microbial metabolism. It is particularly useful for detection of organisms that do not produce conductive metabolites and for samples or media with a high salt concentration.

Method

1. Weigh 0.35g of potassium hydroxide and dissolve in 50ml of deionised water. Do not heat.

2. Weigh 1.0g of Bacteriological Agar No.1 and disperse in 50ml of deionised water in a 100ml screw capped bottle.

3. Fully dissolve the agar, either by boiling or autoclaving.

4. While the molten agar is still hot (approx. 70 ºC), but not boiling, add the cold potassium hydroxide solution and mix thoroughly.

5. Using a suitable pipetting device, dispense 0.7ml of the agar mixture into the base of each clean, dry RABIT cell, taking care to minimize the quantity of agar that runs down the inside of the cell.

6. Allow agar to cool and solidify for between 15 and 30 minutes (maximum), then seal each cell as tightly as possible using the rubber stoppers supplied.

7. After the agar has solidified, indirect cells must be allowed to stabilize for a minimum of 2-3 hours (preferably overnight) prior to use.

8. Indirect RABIT cells may be stored at room temperature for 1 month, if tightly stoppered and protected from strong light. Do not refrigerate the cells.

9. To check the condition of indirect cells before use, place a representative sample in a RABIT incubator module, allow to warm through for 15 to 20 minutes, and observe their conductivity readings using the Global > Conductivity Table function. These values should typically be 7000 - 10000 µS at 30 ºC. Do not use a cell with a conductivity reading below 6000 µS at 30oC.

How do I detect coliforms and E.coli in RABIT?
DETECTION OF COLIFORMS AND
ESCHERICHIA COLI IN RABIT

Introduction

MacConkey Broth is used for the detection of coliforms and E.coli in the RABIT system by the direct impedance technique. Automatic enumeration of coliforms can be achieved by calibrating RABIT results against VRBA pour plates, but it must be noted that RABIT detection of E.coli is on a presumptive basis only.

Preparation

1. Whitley MacConkey Broth is prepared from the dehydrated base at a concentration of 35.0g/l. Sodium deoxycholate must also be added at a level of 0.5g/l to enhance selectivity for the target organisms. Disperse the weighed powders in the appropriate quantity of deionized water.

2. Mix to completely dissolve all the constituents.

3. Dispense the broth into its final containers for autoclaving. If preferred, dispense directly into RABIT cells.

4. Autoclave the broth at 121°C for 15 minutes.

5. Store the prepared broth at room temperature for up to 3 months.

Test Procedure - Coliforms

1. Prepare a tenfold dilution of the product to be tested using a suitable sterile diluent. Maximum Recovery Diluent is recommended for this purpose.

2. Add 1.0ml of sample preparation to 9.0ml of sterile Whitley MacConkey Broth in the direct RABIT cell.
3. Incubate in RABIT using the following test parameters:

Duration : 24h
Time Resolution : 6 min
Detection Criterion : +10 μS
Temperature : 37°C

4. The Time-to-Detection (TTD) generated can be calibrated against a VRBA pour plate count.

5. Lactose fermenting organisms in the test sample are indicated by a change in colour of the broth from purple to yellow.

Test Procedure - E. coli

1. Prepare a tenfold dilution of the product to be tested using a suitable sterile diluent.

2. Add 1.0ml of sample preparation to 9.0ml of sterile McConkey Broth in the direct RABIT cell.

3. Incubate in RABIT using the following test parameters:

Duration : 24h
Time Resolution : 6 min
Detection Criterion : +10 μS
Temperature : 44°C

4. The Time-to-Detection (TTD) generated can be calibrated against a suitable presumptive E. coli plate count method.

5. Presumptive E. coli is indicated by a positive RABIT test result and colour change from purple to yellow in the RABIT cell. However, other thermotolerant coliforms may also produce this result.


How do I detect Yeasts and Moulds in RABIT?

RABIT METHOD FOR THE DETECTION OF YEASTS
AND MOULDS USING SOLID MEDIA


The indirect RABIT method provides a sensitive method for the detection of most yeast species using liquid media. However, detection of moulds is often greatly improved if a solid medium is used, as described below. The indirect method detects microbial growth as a result of carbon dioxide production arising from normal microbial metabolism. It is used for organisms which do not produce conductive metabolites and also for samples or media with a high salt concentration.

Method

1. Suspend an appropriate quantity of Sabouraud Dextrose Agar in deionized water according to manufacturer's instructions and boil to dissolve.

2. Allow to cool to approximately 50°C then pipette 3 ml quantities into each borosilicate glass tube supplied for the indirect RABIT method.

3. Cap the filled tubes (eg Oxoid 12 mm aluminium test tube cap) and autoclave at 121°C for 15 minutes.

4. Remove from autoclave while agar is still molten, place the rack of tubes on an acute slope, and allow to gel.

5. The agar slopes produced in this way should be inoculated with 0.1 ml of sample preparation, which is the optimum volume to coat the surface of the agar.

6. Add the inoculated slopes to prepared indirect RABIT cells and incubate as normal. Typical RABIT parameters would be:

Test Duration 48h
Time Resolution 6 min
Detection criterion -10μS
Temperature 30°C

7. Test duration may need to be increased to 96h and resolution to 12 minutes for slower growing mould strains.


GENERIC RABIT METHOD FOR THE DETECTION OF YEASTS
AND MOULDS USING LIQUID MEDIA

The indirect RABIT method provides a sensitive method for the detection of most yeast species using liquid media. The indirect method detects microbial growth as a result of carbon dioxide production arising from normal microbial metabolism. It is used for organisms which do not produce conductive metabolites and also for samples or media with a high salt concentration.


Culture Media

Choice of culture media will be dependent on sample type and the nature of the expected sample flora. Generally speaking the same media as used in your current conventional methodology should be used.

Choices include Columbia Broth, Wort Broth, Sabouraud Liquid Medium.


Sample Preparation

1. Liquid samples eg water, milk, fruit juice require no preparation although further dilution may be performed as described below. Solid samples require processing to a liquid homogenate at a known dilution.

2. To prepare solid samples: weigh 10 g or 25 g of sample into a sterile Stomacher bag. Add sterile diluent (normally MRD) to ten times the sample weight (eg 100 g, 250 g) and process in the Stomacher until homogenised.

3. Very small samples (eg cosmetics, pharmaceutical preparations) may be diluted by weighing 1 g into a universal bottle containing a measured 9 ml of sterile diluent.

4. If further dilutions of sample are required these should be prepared by pipetting 1 ml of the previous dilution into 9 ml of sterile diluent (normally MRD) and vortex mixing.


RABIT Tests

1. Using the indirect method, culture medium and sample are added to a sterile borosilicate glass tube with a working volume of up to 5 ml. The glass tube is added to a prepared indirect RABIT cell which is then stoppered tightly.

2. The preparation of indirect RABIT cells is described in Technical Note RA5.

3. Each set of RABIT tests must include quality control organisms. These organisms must be grown in pure broth culture then serially diluted in MRD, normally to 10-6. A 1.0 ml aliquot of the highest dilution is then added to the culture medium in a RABIT cell and incubated with the sample cells.

4. Each set of RABIT tests must also include "blank" cells containing sterile culture medium only.

5. After setting up RABIT cells for samples, controls and blanks, cells are incubated in RABIT according to an appropriate Test Code.

6. Typical test codes for general use are shown in the next section. It should be noted that the new “RABIT for Windows” software allows the use of a Total Conductance Change figure as an alternative to the conventional detection criterion.

Typical RABIT Test Codes

                Test                Time                Detection              Temp

                Duration         Resolution     Criterion                (°C)

                (h)                   (min)                (μS)       

 

Test

Code 1     24                6                          -10                          30

 

Test

Code 2    48                 6                          -10                          30

 

Test

Code 3    96                 12                        -15                          30

 

Test

Code 4    24                 6                          -30                          43

                                                   
NOTE: a) By specifying a Total Change threshold as an alternative to the conventional detection criterion, tests run under “RABIT for Windows” software can be made more selective for the target organism(s) by registering equivocal responses as negative.

b) Indirect Test Codes must have a minus value placed in the Detection Criterion.

Enumeration of Yeasts and Moulds

1. For food samples, add 1 ml of sample preparation to 4 ml of Wort Broth in an indirect RABIT cell.

2. For non-food samples, add 1 ml of sample preparation to 4 ml of Sabouraud Liquid Medium in an indirect RABIT cell.

3. If detection of moulds is of the greatest importance, a solid medium should be used, normally Sabouraud Dextrose Agar.

4. Prepare solid medium as slopes in indirect RABIT cell inserts, according to Technical Note RA7.

5. Inoculate slopes with 0.1 ml of sample preparation.

6. Use the following positive control organisms

Wort Broth Saccharomyces cerevisiae
SLM Candida albicans
SDA Aspergillus niger

Control organisms can be sourced from NCTC or ATCC.

7. Incubate all the above media in RABIT according to the appropriate Test Code.

8. The above methods should be compared with a yeast and mould platecount using SDA, Rose Bengal Chloramphenicol Agar or a suitable alternative.

Calibration is achieved by using the RABIT calibration software and entering data for the RABIT TTD and the conventional plate count for a minimum of 30 samples over a range of conventional counts normally encountered in the test sample.

It is critical that the same sample is used to inoculate RABIT and the conventional method when carrying out calibration

How do I clean RABIT cells?
MAINTENANCE AND CLEANING OF RABIT IMPEDANCE CELLS

1. On termination of RABIT tests remove cells from modules and place in suitable racks for autoclaving.

2. INDIRECT CELLS ONLY: Remove bungs from cells and place in a separate container for autoclaving.

3. Autoclave cells/bungs at 121°C for 15 minutes. Do not exceed the specified time or temperature as damage to the cells may result.

4. INDIRECT CELLS ONLY: Remove cells from the autoclave while the agar is still molten. Discard the glass insert tubes and rinse agar from cells under running water. Routine dismantling of cells is not normally required.

5. DIRECT CELLS ONLY: Discard the autoclaved cell contents and separate the cell into its component parts (electrode unit, tube and cap). This should be done every cleaning cycle if cells are heavily soiled by the media/samples used. In cases of lighter soiling full dismantling of cells should take place monthly.

6. Immerse cells/components in a hot 2% solution of Labdet for at least 2 hours, then brush thoroughly to dislodge soiling. Alternatively use the cleaning solution in an ultrasonic bath and ultrasonicate for at least 40 minutes.

7. Rinse cells/components in clean hot water until all traces of detergent are removed.

8. Re-assemble dismantled cells using the cell assembly aid.

9. If cells are to be used for indirect cell production sterilization will not be required.

10. To sterilize cleaned cells autoclave at 121°C for 15 minutes.

11. Dry all cells before re-use by standing in a 40°C cabinet overnight.

12. At least every 5 cleaning cycles, clean the external parts of the cell electrodes by rubbing with a piece of scouring pad in order to maintain a good electrical contact.

Guarantee

The cells supplied with your RABIT system are designed to be durable and robust. Each cell has a one year guarantee provided that the cells are handled in accordance with the manufacturer's guidelines.

NOTE: Use of corrosive substances within the cell could invalidate your guarantee. If in doubt contact Don Whitley Scientific.

What applications does RABIT have within the dairy industry?
IMPEDANCE BIBLIOGRAPHY
DAIRY APPLICATIONS

The following are a selected list of applications of impedance microbiology within the dairy industry. In case of difficulty in obtaining copies of these papers please contact our Technical Support Group.

Asperger H & Pless P (1994)
Salmonella detection in cheese - comparison of methods in regard to the competitive micro-organisms.
Wiener Tierarztlichie Monatsschrift 81, 12-17.

Cousings D L & Marlatt F (1990)
An evaluation of a conductance method for the enumeration of Enterobacteriaceae in milk.
Journal of Food Protection 53, 568-570.

Dziadkowiec D, Mansfield L P & Forsythe S J (1995)
The detection of Salmonella in skimmed milk powder enrichments using conventional methods and immunomagnetic separations.
Letters in Applied Microbiology 20, 361-364.

Firstenberg-Eden R & Tricarico M K (1983)
Impedimetric determination of total, mesophilic and psychrotrophic counts in raw milk. Journal of Food Science 48, 1750-1754.

Gnan S & Luedecke L O (1982)
Impedance measurements in raw milk as an alternative to the standard plate count. Journal of Food Protection 45, 4-7.

Hancock I, Bointon B M & McAthey P (1993)
Rapid detection of Listeria species by selective impedimetric assay.
Letters in Applied Microbiology 16, 311-314.

Joosten H M L J, Van Der Velde V D & Van der Velde F (1994)
Evaluation of motility enrichment on modified semisolid RV medium and automated conductance in combination with Rambach agar for Salmonella detection in environmental samples from milk powder.
International Journal of Food Microbiology 22, 201-206.

Kamei T, Sato J, Kodama Y, Omata Y & Noda K (1988)
Application of the conductance method to detection of post-pasteurization contamination of pasteurized milk.
Nippon Shokuhin Kogyo Gakkaishi 35, 226-234.

Lanzanova M, Mucchetti G & Neviani E (1993)
Analysis of conductance changes as a growth index of lactic acid bacteria in milk. Journal of Dairy Science 76, 20-28.

Madden R H & Gilmour A (1995)
Impedance as an alternative to MPN enumeration of coliforms in pasteurised milks. Letters in Applied Microbiology 21, 387-388.

McPhillips J & Snow N (1958)
Studies on milk with a new type of conductivity cell.
Australian Journal of Dairy Technology 3, 192-196.

Mosteller T M & Bishop J R (1993)
Sanitizer efficacy against attached bacteria in a milk biofilm.
Journal of Food Protection 56, 34-41.

Neaves P, Waddell M J & Prentice P J (1988)
A medium for the detection of Lancefield Group D cocci in skimmed milk powder by measurement of conductivity changes.
Journal of Applied Bacteriology 65, 437-448.

Nieuwenhof F F J & Hoolwerf J D (1987)
Impedimetric detection of post-pasteurisation contamination on pasteurised milk. Netherlands Milk Dairy Journal 41, 49-68.

O’Connor F (1979)
An impedance method for the determination of bacteriological quality of raw milk. Irish Journal of Food Science and Technology 3, 93-100.

Phillips J D & Griffiths M W (1989)
An electrical method for detecting Listeria spp.
Letters in Applied Microbiology 9, 129-132.

Pirovano F, Piazza I, Brambilla F & Sozzi T (1995)
Impedimetric method for selective enumeration of specific yoghurt bacteria with milk-based culture media.
Lait, 75, 285-293.

Prentice G A, Jervis D I, Ester M C & Neaves P (1990)
An interlaboratory evaluation of an electrometric method for detection of Salmonellas in milk powders (Rapid Microbiological Methods for Foods, Beverages and Pharmaceuticals).
SAB Technical Series, 155-164

Silley P & Forsythe S (1996)
Impedance microbiology - a rapid change for microbiologists.
Journal of Applied Bacteriology 80, 233-243

Suhren G & Heeschen W (1987)
Impedance assays and the bacteriological testing of milk and milk products. Milchuissenschatt Milk Science International 42, 619-627

Svensson U K (1994)
Conductimetric analysis of bacteriophage infection of 2 groups of bacteria in DL-lactococcal starter cultures.
Journal of Dairy Science 77, 3524-3531.

Taranto M P, Hogado P de R & Valdez G F de (1997)
Use of a conductimetric method to evaluate the effect of bile acids on Listeria monocytogenes.
Milchwissenschaft 52, 247-249.

Visser I J R & de Groote J M F H (1984)
The Malthus microbiological growth analyser as an aid in the detection of post-pasteurization contamination of pasteurized milk.
Netherlands Milk Dairy Journal 38, 151-156.

Visser I J R & de Groote J M F H (1984)
Prospects for the use of conductivity as an aid in the Bacteriological Monitoring of Pasteurised Milk.
Antoine van Leewenhoek 50, 202-206.

Yoshida K, Haruta M, Kitada T, Suzuki I & Norichi T (1987)
A rapid estimation of the number of lactic acid bacteria in fermented milk and lactic beverages via the conductance measurement method.
Japanese Journal of Zootechnical Science 58, 13-20.


Can I use RABIT for biofilm evaluation?
RABIT APPLICATIONS - BIOFILMS

There are a number of publications where impedance has been used as a tool in the study of biofilms. Published papers refer to the use of RABIT and other competitor instruments specifically Malthus and Bactometer.

Holah et al (1990) described the use of direct impedance to enumerate microbes on steel discs following exposure to 12 disinfectants. The ‘pass’ criterion was a 5 log reduction in test organism viability after 5 min exposure. Similarly, Druggan et al (1993) used indirect impedance with 1 cm diameter steel discs which were directly transferred to the impedance tubes. This avoided the inaccuracy of physical removal of the microbial growth using sand agitation. Sodium hypochlorite was used as the model for chlorine-based disinfectants. The European suspension test organisms Ps.aeruginosa NCIB 10421, Proteus mirabilis NCIB 12596, Staph.aureus NCTC 10788 and S.cerevisiae ATCC 9763 were used to test the efficacy of sodium hypochlorite to microbial biofilms. Biofilms were produced by the three bacterial strains but not by the yeast. The detection time of the bacteria biofilms did not correspond with the cell density possibly due to differences in microbial metabolism rates. For example Ps.aeruginosa produced a biofilm of 5.0 x 106 cfu disc which gave a detection time of 4.0 h in WIB, whereas Staph.aureus biofilm was 2.0 x 107 cfu per disc with a detection time of 6.5 h. Johnston and Jones (1995) used a Modified Robbins Device (whereby a microbial culture is circulated over steel discs) to produce biofilms of Ps.aeruginosa. Enumeration by indirect impedance showed higher numbers of surviving cells than cell recovery by swabbing or vortexing.

Flint et al (1997) used impedance to study the numbers of thermophilic streptococci on stainless steel and Andrade et al (1998) showed that impedance was far superior to standard plate counts for assessing the effect of sanitizers on Enterococcus faecium biofilm attached to stainless steel surfaces.

Mosteller and Bishop (1993) showed that Ps.fluorescens, Y.enterocolitica and L.monocytogenes readily attach to rubber and Teflon® surfaces. The test organisms attached in slightly higher numbers to the rubber surface than the Teflon®. Plate counts, impedance microbiology and the direct epifluorescent filter technique were compared. Impedance microbiology was the best method of enumeration since it allowed the estimation of both reversibly and irreversibly attached bacteria. Biocides against a bacterial suspension resulted in a greater than or equal to 5 log cycle reduction. However, the same concentrations were relatively ineffective against the attached bacteria. The goal reduction (3 log cycles) was achieved on the Teflon® surface with the iodophor, hypochlorite and fatty acid biocides with a log-cycle reduction in the number of Y.enterocolitica of 3.09, 3.19 and 3.31 respectively. Pseudomonas fluorescens was reduced by 3.16 on both the rubber and Teflon® surfaces when exposed to the hypochlorite biocide.

Microbially influenced corrosion affects various industries but can be partially controlled by the application of biocides. Copper surfaces exposed to natural seawater were colonized by bacteria within 3 weeks of exposure independent of alloy composition (Mansfeld and Little 1992). Jack et al (1992) and Nivens et al (1992) studies the corrosion rates of carbon steel by monocultures and various combinations of Bacillus sp, Hafnia alvei and Desulfovibrio gigas biofilms in an aerobic, continuously flowing freshwater reactor containing 0.4 mmol 1-1 sulphate. Debruyn et al (1994) correlated the viable count of D.desulfuricans in iron sulphite medium with impedance microbiology (r = 0.974). Subsequently the impedance method was used to assess the efficacy of biocides against D.desulfuricans. A 56% and a 100% kill was obtained using 60 and 200 mg 1-1 quaternary ammonium compounds respectively.

References

Andrade N J, Bridgeman T A and Zottola E A (1998)
Bacteriocidal activity of sanitizers against Enterococcus faecium attached to stainless steel as determined by plate count and impedance methods.
Journal of Food Protection 61, 833-838.

Debruyn E E, Croukamp E and Cloete T E (1994)
The Malthus system for biocide efficacy testing against Desulfovibrio desulfuricans. Water SA 20, 23-26.

Druggan P, Forsythe S J and Silley P (1993)
Indirect impedance for microbial screening in the food and beverage industries.
In New Techniques in Food and Beverage Microbiology ed Kroll R G, Gilmour A and Sussman M. Society for Applied Bacteriology, Technical Series No 31. Oxford. Blackwell Science.

Flint S H, Brooks J D & Bremer P J (1997)
Use of the Malthus conductance growth analyser to determine numbers of thermophilic streptococci on stainless steel.
Journal of Applied Microbiology 83, 335-339.

Holah J T, Higgs C, Robinson S, Worthington D and Spenceley H (1990)
A conductance-based surface disinfection test for food hygiene.
Letters in Applied Microbiology 11, 255-259.

Jack R F, Ringelberg D B and White D C (1992)
Differential corrosion rates of carbon-steel by combinations of Bacillus sp., Hafnia alvei and Desulfovibrio gigas established by phospholipid analysis of electrode biofilm. Corrosion Science 33, 1843-1853.

Johnston M D and Jones M V (1995)
Disinfection tests with intact biofilms - combined use of the Robbins device with impedance detection.
Journal of Microbiological Methods 21, 15-26.

Mansfeld F and Little B (1992)
Microbiologically influenced corrosion of copper-based materials exposed to natural seawater.
Electrochimica Acta 37, 2291-2297.

Mosteller T M and Bishop J R (1993)
Sanitizer efficacy against attached bacteria in a milk biofilm.
Journal of Food Protection 56, 34-41.

Nivens D E, Jack R, Vass A, Guckert J B, Chambers J Q and White D C (1992) Multielectrode probe for statistical evaluation of microbiologically influenced corrosion. Journal of Microbiological Methods 16, 47-58

In addition to the published studies DWS have a number of customers currently using RABIT to study biofilm development most notably in the oral and health care sectors.

Can RABIT detect thermophillic organisms?
DETECTION OF THERMOPHILIC ORGANISMS IN RABIT

Two detection methods are available to RABIT users, the direct and indirect methods. The indirect method does not work particularly well at elevated temperatures due to water vapour condensing at the top of cell and dropping back onto the potassium hydroxide bridging plug thereby altering the conductivity of the agar plug and giving rise to a “noisy” electrical signal.

As a general principle RABIT modules can be run up to a temperature of 50°C using the direct impedance method. DWS have experience of working with Whitley Impedance Broth and Whitley Anaerobe Broth at elevated temperatures and in each case there can be problems caused by evaporation of the culture medium and an associated change in medium conductivity. This can be reduced by applying a layer of sterile liquid paraffin or mineral oil to the surface of the culture medium after sample inoculation. This will obviously restrict oxygen supply to the inoculated culture and is thus primarily targeted at growth of thermophilic anaerobes.


How do I work out generation times of organisms using RABIT?

DETERMINATION OF MICROBIAL GENERATION TIMES USING RABIT


Generation time of a microbial cell is the time taken for one cell to divide into two cells. It is also referred to as doubling time.

Conventionally this parameter is determined from classical growth curves where numbers of organisms in a culture are counted over a time course and the corresponding generation time is determined from a graph of colony forming units against time.

As Impedance microbiology is simply a measure of microbial growth it can be readily used to determine generation or doubling time without the need to construct growth curves from plate count data.


Principle

If a series of serial dilutions are prepared from an initial microbial culture then the generation time (tg) can be calculated by determining the delay in time to detection (TTD) between the respective dilutions.

Using a known dilution factor, normally 1:100, it becomes unnecessary to even know the exact concentration of organisms in either dilution.

Procedure

Using dilutions with a hundred fold difference then the formula

tg = is used

TTD = the differences in TTD between the respective dilutions used in the calculation

Example:

Inoculate the test culture into Whitley Impedance Broth and incubate overnight.

Prepare a set of serial dilutions and for the RABIT test use the 10-3 and the 10-5 dilution. Thus you have a hundred fold difference between the two dilutions.

Take 1 ml of each of the above dilutions and inoculate into 9 ml WIB and set up test in RABIT to determine TTD.

Assume TTD of 10-3 dilution is 8h 30 minutes
and TTD of 10-5 dilution is 12 h 30 minutes

Calculation:

tg =

NB - log2 is used because we had a 100 fold difference (log2) between the two dilutions

` So tg =

= 0.602 hours

= 36.1 minutes


The formula can be simplified whenever a hundred fold dilution is used because log n1 - log n2 will always be 2 and as log2 = 0.301 the formula becomes

tg = =

= 0.15 TTD

In the example above

tg = 0.15 × 4 = 0.6 hours = 36.1 minutes

How does impedance compare with ATP technology?
IMPEDANCE VERSUS ATP TECHNOLOGY
IS THERE A DIFFERENCE?

Impedance and ATP technologies have both been with us for more than 20 years as viable and useful technologies. In that time they have been refined and protocols have been developed which have clearly shown both technologies to be relevant to the current demands of a busy microbiology laboratory.

ATP based systems are now largely used within the hygiene monitoring sector and in this mode they are highly successful providing a rapid result usually within 30 minutes and often in a much shorter time frame, depending on the system and the application. Impedance, however, has remained targeted at the quality assurance and research and development sectors with results being related to quantitative enumeration of microbial activity. Whilst results can be available within two hours for samples with a high bioburden they may take up to 24 hours for samples with a low total count, yet in all cases they will be available more quickly than for routine cultural approaches.


Technology Difference

There is a fundamental difference between the two technologies with regard to the limit of sensitivity. Impedance will detect down to 1 viable cell within the test sample. If 1 viable cell is present in the sample and the growth medium in the impedance test cell is capable of supporting the growth of that 1 organism then impedance will detect its presence assuming the test is run for a sufficient time to enable growth to occur. The lower limit of detection is thus 1 viable cell.


By contrast ATP technology is dependent upon the number of cells present at the time the sample is taken. The number of cells present when the sample is taken must be higher than the detection threshold. If we assume the limit of detection to be 103 cfu/ml then a positive result will only be detected if there are greater than 103 cfu/ml at the point the sample is taken. If at this time there were 8 x 102 cfu/ml then the test would appear to be negative.

It is clear that impedance is inherently more sensitive than ATP-based technology.

What is the best way to check inhouse if the RABIT system is performing correctly?
RABIT TEST CELLS

A precision set of high quality test cells for use exclusively with the RABIT system produced by Don Whitley Scientific Limited. Housed in an attractive mahogany presentation box and complete with instructions for use.

Each set comprises:

Temperature Test Cells
3 ´ Test Cells covering the temperature range 20°C - 50°C.
Each Temperature Test Cell comprises a high quality thermometer protected by a clear Pyrex glass tube which permits the thermometer scale to be read. The base plug of the cell is precision machined from brass for good thermal conductivity and is finished in a hard chrome coating for durability. The top plug is precision machined from aluminium and protected by clear anodising. Each top plug carries an identification label, sub-surface printed for wear resistance.

Impedance Test Cells
5 ´ Test Cells covering the range 4000 µS - 20000 µS.
Each Impedance Test Cell is capable of measuring a designated impedance value. The five individual cells are: 4000 µS; 8000 µS; 12000 µS; 16000 µS and 20000 µS. The central portion of each Test Cell is polypropylene, sheathed internally with a grey PVC liner to protect the electronic components. The lower portion is a standard RABIT cell plug which enables an electrical connection to be made to the module. The top plug is precision machined from aluminium and protected by clear anodising. Each top plug carries an identification label, sub-surface printed for wear resistance.

What levels of microbes do I need in my samples in order to use a spiral plater?
UPPER AND LOWER COUNT LIMITS FOR SPIRAL PLATES


In order to explain the count limits it is first necessary to understand the method of counting for spiral plates.

All spiral plates are prepared in an identical manner whereby a fixed volume of sample is deposited on the surface of an agar plate at a decreasing rate from the centre to the edge, therefore fixing the volume/area relationship for any portion of the plate. In a mode where 50 µl is dispensed a dilution of 1000:1 is created across the plate which will produce a countable plate from any sample containing between 400 cfu/ml up to 4 x 105 cfu/ml without the need for further dilution.

After incubation colonies appear along the track made by the deposited sample, with spacing between colonies increasing from the centre to the edge.

The viewing grid for counting spiral plates is shown in Fig 1.

The major divisions on this grid are the concentric circles and the 8 pie-shaped wedges or sectors which result in a number of annular segments.


Counting is done by placing the petri-dish over the grid and counting along the pie-shaped wedge - from the outside towards the centre - until at least 20 colonies are counted, then completing the count in the segment in which the 20th colony was found. The opposite segment is then counted.  The sum of these 2 counts, divided by the known segment pair volume, gives the concentration.

Why there is a minimum count rule of 20 can be seen by looking at the percentage co-efficient of variation (CV) when applied to plate counts. This is the standard deviation of replicate counts from separate plates divided by the mean counts × 100. The following table shows CV for spiral plate counts in 3 ranges: 0-5, 6-20 and 21-200 cfu.


    Colonies counted per plate    
    (CV %)       

     0 - 5 (78)
     6 - 20 (25)
    21 - 200 (15)
    

Comparing these figures it can be seen that until the number of colonies counted reaches 20, the accuracy of the count is low. Above 20, accuracy compares favourably with the standard plate count.

An upper limit of 75 colonies in each sector is set as above this number the accuracy of the count will be low due to coincidence error associated with the crowding of colonies. However to count 75 colonies in a sector assumes regular sized 1-2 mm colonies and would not apply with large irregular colonies which will overlap at lower densities.


How do I prepare agar plates for use with my spiral plater?
PREPARATION OF AGAR PLATES - SPIRAL PLATING

Repetitive and predictable deposition of a solution on the surface of a prepared agar plate is a primary requirement of spiral plating. Although the Spiral Plater, when properly adjusted, will precisely deliver the volume from the stylus tip the amount and distribution of the sample on the surface of the agar surface is influenced by the condition and character of the agar surface itself. This is especially apparent on areas of the plate where the volume dispensed is very low and where factors other than the displacement of fluid by the syringe become operative, such as agar drag against the stylus tip. It is necessary that the agar surface be smooth without wrinkles, pits, or air bubbles; that the agar be of uniform depth over the entire plate; that no free water droplets are on the surface; that there are no areas of dehydration; and that there is no contamination on the plate from bacteria or mould.

The following remarks on the consequence of various agar defects provide the reasons for concern and provide a rationale for care in preparation.

If air bubbles are present when pouring agar into the plates, they break upon solidification and leave pinholes or cavities which receive additional fluid from the stylus as it moves over the agar surface.

The same quantity of agar should be poured into all plates in order that the same height of agar will be presented to the stylus tip. The agar plates should be level when cooling, resulting in the same depth of agar in the Petri dish. The Spiral Plater will accommodate some unevenness in depth but obvious unevenness will result in a larger quantity of sample being deposited in one side of a plate.

Drops of water on the agar surface will flow, and wash, or move, bacteria from their deposited position. While the colony pattern resulting from this movement is readily seen and though usually colonies on some portions of a plate can be counted, it is not a good practice.

Prepared agar plates which have been dried excessively tend to grow fewer colonies on the outer portions of a spiral plate than “normal” plates.   Another problem is encountered when plates are cold at the time of plating. Liquid from the agar wells up along the track made by the stylus tip on the agar surface. This liquid is not quickly reabsorbed and tends to flow across tracks, thus destroying the spiral pattern normally made.

For any comparison of plating procedures, the same brand and lot number of nutrient agar should be used. For spiral plating, treat the prepared plates with a standard procedure of preparation, storing, tempering, and use. A suggested procedure is as follows:

a)    Pour the sterile agar at about 45-50°C.

b)    Put a standard amount of agar into each dish -about 45-50 ml into 15 cm Petri dishes and about 20 ml into 10 cm Petri dishes.

c)    Allow the agar to solidify on a level surface with the poured plates stacked no higher than 10 to a stack.

d)    Before use plates should be dried in a drying oven at a temperature not greater than 55°C, for 12 minutes, or in a 37°C incubator for 1 hour, or until all excess water has evaporated from the surface of the agar.


How can I be sure the stylus on my WASP is being cleaned efficiently?

WASP CLEANING EFFICIENCY

1.    INTRODUCTION

Don Whitley Scientific recommend the use of sodium hypochlorite or ethanol as sanitising fluids for the WASP spiral plater. The listed sanitisers have a long history of use with respect to spiral plating and indeed were used in the initial validation studies in which there were no reports of cross contamination between samples as a result of the spiral plating technique.

In order to build upon a proven technology DWS utilised the same teflon material in the WASP stylus assembly as in the original platers. The combination of teflon tubing, induced turbulence within the tubing in the wash cycle and an effective sanitiser ensure that the WASP stylus assembly is adequately decontaminated between samples. This claim has been fully substantiated within the DWS laboratories as indicated by the following data.

Since the studies described in this Technical Note were completed it has come to our attention that in some instances repeated use of oily products such as cream (dairy and non-dairy) can cause a build up of residual sample material within the stylus. In these cases we recommend that a warm 1% solution of Labdet be used to flush the stylus assembly.


2.    EXPERIMENTAL PROTOCOL

2.1    Test Organisms

Bacillus subtilis globigii, an orange pigmented colony
Salmonella typhimurium, white pigmented colony

2.2    Procedure

2.2.1    4 x 50 µl aliquots of B.subtilis globigii plated onto Plate Count Agar (PCA) after which WASP was sanitised using sodium hypochlorite and the automatic wash procedure.

2.2.2    Following washing 4 x 50 µl aliquots of sterile water were plated onto PCA followed again by the automatic sanitise/wash routine.

2.2.3    Step 2.2.1 was repeated but with Salmonella typhimurium.

2.2.4    At the end of the study the sterile water wash pots were plated onto PCA.

2.3    Results

2.3.1    Orange pigmented colonies of B.subtilis globigii were present on PCA following 2.2.1.

2.3.2    No growth was detected on PCA from 2.2.2 clearly showing the sanitisation following inoculation of 4 x 50 µl of B.subtilis globigii had been complete.

2.3.3    White pigmented colonies of S.typhimurium were present on PCA following 2.2.3.   The absence of any orange pigmented colonies clearly showed there was no residual carry over from 2.2.1.

2.3.4    No growth was determined following plating of the sterile water wash pots at the end of the study, showing that sanitisation was complete and that the wash cycle purely served to remove any residual sanitiser from the stylus tubing.

2.4    Repeat Study with Oily Product

2.4.1    The initial study described above was carried out with pure cultures. In order to simulate a worst case scenario soft white paraffin was seeded with B.subtilis globigii or with S.typhimurium and the above study repeated.

2.5    Results

Results were as described in the initial study.

3.    CONCLUSIONS

Sanitisation with sodium hypochlorite is fully effective and prevents cross contamination between samples whether working with aqueous or oily based products.

What daily checks should I carry out on my Spiral Plater?
RECOMMENDED DAILY CHECK PROCEDURE
    FOR WHITLEY AUTOMATIC SPIRAL PLATER (WASP)


Don Whitley Scientific Limited recommend that the following daily checks are performed on the Whitley Automatic Spiral Plater prior to use to ensure correct functioning.

The following is a summary of these checks.   For full details please refer to the instrument’s User Manual.

1.    Check that the level of vacuum available is between 18" - 22" Hg.

2.    Check that liquid passes quickly through the sight glass when the stylus is out of the wash pots during the washing sequence.

3.    Fill a dispo beaker with sterile water and draw up the entire contents whilst monitoring the content passing through the sight glass. Ensure no bubbles are seen.

4.    Check that the petri dishes being used fit the turntable and are centred correctly.

5.    Check the condition of the stylus, it should be free from physical damage, blockage and kinks.

6.    Perform a dye plate using water soluble ink. Whilst the system fills with ink check that no air bubbles are visible in the sight glass. The track produced should be uninterrupted with no surface skipping.

7.    Initiate the test routine to ensure the stylus tip start and finish points are correctly set.


8.    Check the batch of agar plates being used. A good agar plate for spiral plating should have:-

a.    A smooth surface with no pits or airbubbles.
b.    A uniform depth of agar over the entire plate.
c.    No water droplets on the surface.
d.    No areas of dehydration.
e.    No microbial contaminants.

For further details refer to Technical Note WA6 ‘Preparation of agar plates - spiral plating’.

How accurate is my spiral plater compared to my existing method?

SENSITIVITY AND REPRODUCIBILITY OF WASP -

INTRODUCTION

Traditional methods for counting viable microorganisms are labour intensive yet the standard plate count remains a widely used tool in many microbiology laboratories. The spiral plate method for the enumeration of organisms introduced considerable advantages over the standard plate count and has been shown to give comparable results. The technique was, however, limited in that it did not allow deposition of replicate samples without refilling the syringe and more importantly it had a lower sensitivity of 400 cfu/ml as a result of the 500 µl sample volume.

METHODS
 
All studies were carried out on a production WASP instrument. The WASP uses microprocessor-controlled stepper motors to automate the instrument and consequently allows the sample volume to be selected as 50, 100, 200 or 400 µl.

Determination of dispensed volume

Determination of dispensed volume was in accordance with BS7653:  Part 3:  1993, Piston and/or plunger operated volumetric apparatus:  Methods of test. A distilled water sample was loaded by the instrument in either the 50, 100 or 200 µl mode and then dispensed and collected into a clean, dry, pre-weighed flask which was immediately sealed after all sample had been dispensed. For 50 µl sample volumes this procedure was repeated four times into the same flask. The flask was then re-weighed on an analytical balance and the average weight for each aliquot determined.

Determination of lower sensitivity of WASP

Escherichia coli (NCTC 10418) was used for all studies. An overnight broth culture (Tryptone Soya Broth) of the test organism, incubated at 37°C was diluted to give a series of suspensions with nominal concentrations in the range 10-1000 cfu/ml sample. From each of these dilutions 1 ml pour plates and spiral plates in which WASP was set to dispense 200 µl sample, were inoculated onto Plate Count Agar (PCA) and incubated at 37°C overnight. Five replicate plates were inoculated at each dilution and for each method. All the colonies were counted on the pour and spiral plates.
Reproducibility of plate counts

The method was as described for determining the lower sensitivity of WASP except that the overnight broth culture was diluted to give counts in the range 1000-10000 cfu/ml. From each of the dilutions spiral plates were inoculated onto PCA using the 50, 100 and 200 µl deposition modes. Appropriate dilutions were made and 1 ml pour plates also inoculated onto PCA. Five replicate plates were inoculated for each dilution and colonies were counted after overnight incubation at 37°C. Spiral plates were counted according to the normal 20/20 rule whereby the agar plate is laid over a counting grid and at least 20 colonies are counted in a specified grid sector and in the equivalent opposite sector, this being used to relate the number of colonies in that sector to the total sample volume dispensed within that same sector.

Determination of microbial loading in ice cream

A 25 g aliquot of dairy ice cream was added to 225 ml MRD and stomached for 30 seconds. Five replicate samples were inoculated onto PCA using the WASP spiral plater for each of the three deposition volumes 50, 100 and 200 µl. Plates were incubated overnight at 30°C and all colonies counted manually.

TEST RESULTS

Table 1        Reproducibility of counts from the standard pour plate method and WASP using 50 µl, 100 µl and 200 µl deposition modes

 
     WASP Count (cfu/ml)       
  50 µl deposition    100 µl deposition    200 µl deposition       
    Mean    SEM         Mean    SEM          Mean    SEM          Mean    SEM       
1.1 ´ 104    706    1.2 ´ 104    510    1.1 ´ 104      245    1.4 ´ 104    1,400       
5.9 ´ 103    288    7.7 ´ 103    759    6.2 ´ 103      339    6.3 ´ 103       383       
4.0 ´ 103    283    4.6 ´ 103    615    7.2 ´ 103    1,714   3.8 ´ 103       153       
3.2 ´ 103    258    3.1 ´ 103    103    4.3 ´ 103      255    4.8 ´ 103       103       
2.1 ´ 103    308    2.4 ´ 103    114    3.3 ´ 103      480    2.9 ´ 103       132       
1.1 ´ 103    105    1.0 ´ 103      43    9.8 ´ 102        55     1.1 ´ 103        65    


Table 2        Reproducibility of the dispensed volume of the WASP in 50 µl, 100 µl and 200 µl mode

 
                             & nbsp;                         &n bsp;             50 µl       100 µl             200 µl       
Number of Observations (n)                  24              12                    12       
Range of Dispensed Volume (µl)    50-51    101.0-102.1    197.0-204.9       
Mean Volume Dispensed (µl)            50.7            101.7             201.6       
Standard Error of the Mean (SEM)     0.04             0.08               0.73    

Table 3        Determination of the lower sensitivity level of WASP in the 200 µl deposition mode

 
                             & nbsp;                         &n bsp;           n    cfu/ml    SEM       
Standard Pour Plate Count (cfu/ml)    5      31         3.2       
WASP Count (cfu/ml)                              5     30          5.2    


Table 4        Reproducibility of total plate counts from ice cream using WASP with 50, 100 and 200 µl deposition modes

 
Mode                             & nbsp;       50 µl    100 µl    200 µl       
Mean count per plate          7.8       15.0       29.2       
cfu/ml                             & nbsp;       156       150        146       
CV1                        &nbs p;                45.6     26.3       14.4    

1    Coefficient of Variation (Standard Deviation ´ 100/mean value)


DISCUSSION

The basis of the spiral plating technique is that a known volume of sample is dispensed onto a rotating agar plate in an Archimedes spiral. The volume of sample deposited per unit area of plate decreases across the spiral resulting in a dilution effect, and allowing counting of isolated colonies in respective sectors of known deposition volume. The original spiral plating instruments all utilised a mechanical system to drive sample deposition whereby a cam follower linked to the syringe moved along a pre-defined path to determine the total volume and profile of the dispensed sample. The mechanical system has now been replaced by software and electronic stepper motors such that the instrument may be automated.   The dispensed volume is still crucial to the final calculation and the test result. The data clearly show the reproducibility of the WASP whether in the 50, 100 or 200 µl mode. The ability to demonstrate delivery of stated volumes is particularly relevant to those laboratories seeking accreditation. The requirement to count a minimum of 20 colonies on an agar plate using the spiral plate technique coupled with a sample volume of 50 µl, ensured that the lower limit of counting for the original mechanical platers was 400 cfu/ml. The new generation of automated spiral platers have the facility to deposit 200 µl aliquots and for samples in which the microbial loading is expected to be very low it is appropriate to count the total number of colonies on the plate, rather than utilising a sector count.   The lower limit in this study was 30 cfu/ml at which point it was equivalent to the count from a standard pour plate with 1 ml sample. The data presented substantiates earlier studies showing the spiral plating technique to be equivalent to the pour plate method. Subjecting the data to one way of analysis of variance showed there to be no significant differences between the data sets from the 50, 100 or 200 µl deposition format. These findings were further corroborated following analysis of the data from the ice cream counts.

CONCLUSIONS

The results of this study show that data generated by the new generation of automated spiral plater are equivalent to the standard pour plate count and that by virtue of the increased deposition volumes the technique now shows an increased sensitivity, such that microbial loadings as low as 30 cfu/ml can be accurately plated and counted. The data from the ice cream clearly show the benefits of being able to inoculate a larger sample volume onto a spiral plate in that the CV reduces from 45.6% for a 50 µl sample to 14.4% for the 200 µl sample thereby increasing confidence in the test data. This is particularly relevant with respect to due diligence within the food industry.

How fast will the spiral plater process a sample?
WASP TIMINGS

Introduction

WASP spiral platers are subject to continued development and consequently it is necessary to periodically report on the current specifications of the plater.   This technical note details the times taken for basic operations.

Timings

1.    A basic WASH cycle takes 19.3 seconds.

2.    A single 50 µl log plate including a 5 second sample uptake time takes 19.7 seconds.

3.    A single 50 µl log plate followed by a wash cycle (user initiated) takes 39 seconds.

4.    A second and subsequent 50 µl log plate takes 10.8 seconds.

5.    For example 4 ´ 50 µl plates followed by a wash cycle would take

19.7 + 10.8 + 10.8 + 10.8 + 19.3 = 71.4 seconds

How is the spiral plater checked to ensure it is dispensing the correct volume?
WASP - GRAVIMETRIC ANALYSIS

INTRODUCTION

The volumes to be dispensed by the WASP instrument are factory set, however, many laboratories will want to carry out regular checks with respect to the volume dispensed.

The gravimetric determination of small volumes requires an analytical balance and adherence to strict procedures to ensure reproducibility and accuracy. For further information on this method refer to British Standard BS 7653, Part 4, 1993; Piston and/or plunger operated volumetric apparatus.  Specification for conditions of test, safety and supply.

All equipment, materials, test water and the WASP must be allowed to equilibrate for two hours prior to use in the environment where testing will take place.

PROCEDURE

1.    Turn on both the vacuum source and WASP.

2.    Place a clean, dry, sealable container in which to collect the sample onto an Analytical balance and press the “Tare” key so that the display reads zero.

3    Set the WASP in the following mode:-

Auto/Valve/Log/50µl

Open the valve on the WASP and take up a sample of deionized water.   press the “start plating” key and, using the sealable container collect the dispensed water.   Keep the water in a single droplet and avoid warming the container with the hand to avoid evaporation.

4.    Repeat a further three times, replacing the lid of the container between depositions.  Place the container onto the balance and record the reading, to four decimal places immediately.   Divide this reading by four to obtain the mean deposition value.   Repeat four times and calculate the overall mean deposition value.

5.    Place a further clean, dry, sealable container onto the analytical balance and tare the balance.


Set the WASP in the following mode:-

Manual/Syringe/Log/50µl

Fill the syringe of the WASP with a sample of deionized water.   Press the “start plating” key and, using the sealable container collect the dispensed water as 3.   Repeat this a further 3 times replacing the lid of the container between depositions.   Place the container onto the balance and record the reading to four decimal places immediately.   Divide this reading by four to obtain the mean deposition value.   Repeat four times and calculate the overall mean deposition value.

6.    Place a further clean, dry, sealable container onto the analytical balance and tare the balance.

Set the WASP in the following mode:-

Auto/Valve/Log/100µl

Open the valve and take up a sample of deionized water.   Press the “start plating” key and, using the sealable container collect the dispensed water as in 3.   Place the container onto the balance and record the reading to four decimal places immediately.  Repeat four times.

7.    Place a further clean, dry, sealable container onto the analytical balance and tare the balance.

Set the WASP in the following mode:-

Auto/Valve/Log/200 µl

Open the valve and take up a sample of deionized water.   Press the “start plating” key, and, using the sealable container collect the dispensed water as in 3.   Place the container onto the balance and record the reading to four decimal places immediately.  Repeat four times.

INTERPRETATION OF RESULTS

When the WASP is set in the mode whereby 50 µl is deposited onto the agar a volume of 50 ± 2 µl must be obtained for each deposition.

Both 100 µl and 200 µl depositions are absolute multiples of the 50 µl deposition therefore a volume of 100 ± 4 µl must be obtained when the WASP is set to dispense 100 µl a volume of 200 ± 8 µl must be obtained when the WASP is set to dispense 200 µl.


What accreditation does the WASP carry?
SPIRAL PLATING APPROVAL


The spiral plate count method for microorganisms in milk, foods and cosmetics is an official method of the American Public Health Association (APHA), the Association of Official Analytical Chemists (AOAC) and the British Standards Institute (BSI).

Detailed reference to these methods can be provided.


References:

1.    American Public Health Association (1984) Compendium of Methods for the Microbiological Examination of Foods, 2nd Ed. APHA, Washington DC

2.    Association of Official Analytical Chemists (1990) Official Methods of Analysis, 15th ed. AOAC, Arlington, VA, USA

3.    Microbiological examination for dairy purpose. Part 2.  Methods of general application for enumeration of microorganisms.   Section 2.3 Enumeration of microorganisms by surface plate technique for colony count.

4.    AFNOR (France) Spiral Plating is recognised as standard VO8-100 under programme no 59 of the AFNOR accreditation for food microbiology. The methods for TVC and total coliforms have been accredited under this programme.

How do I make the sanitising solution for the WASP?
DISINFECTANT (SANITIZER) SOLUTIONS FOR USE WITH WHITLEY AUTOMATIC SPIRAL PLATER (WASP)

Don Whitley Scientific Limited has evaluated three categories of disinfectant agent for use with the WASP instrument. It is essential that WASP is used with one of the recommended agents at the correct concentration:  this will avoid invalidation of the instrument warranty for WASP and any accompanying vacuum source.

1.    Sodium hypochlorite solution (bleach)

Commercial sodium hypochlorite solutions used to prepare WASP disinfectant should contain an available chlorine concentration of between 10% and 15% (100,000 ppm to 150,000 ppm) as supplied. For use with WASP, such solutions must first be diluted in water to 5% of the original concentration. This will produce a final available chlorine concentration of 5000 ppm (nominal). If the concentrate is purchased at a different strength then the relative proportions should be adjusted accordingly.


2.    Chlorine Release Tablets

DWS has evaluated commercially available chlorine release tablets/effervescent disinfectant tablets containing sodium dichloroisocyanurate (NaDCC). Suitable products include Preseptä and Haztabsä. If such a product is used with WASP, it must first be fully dissolved in water to give a final available chlorine concentration of between 2000 ppm and 2500 ppm. This concentration range is typically recommended for use in laboratory “discard jars” (eg for pipette disinfection).

3.    Ethanol

Industrial ethanol (industrial methylated spirit) may be used with WASP at a concentration of 70% v/v in water. It is essential that a 70% v/v solution is used, to ensure maximum disinfectant efficacy.

NOTE: Studies at DWS have indicated that the disinfectant efficacy of ethanol may be less than that of sodium hypochlorite or chlorine release tablets.

UNSUITABLE AGENTS: Calcium hypochlorite (bleaching powder) must not be used in WASP.   Use of this agent will invalidate the instrument warranty.

How do I prevent the stylus on my WASP blocking?

WASP – PREVENTION OF STYLUS BLOCKING


From time to time it is possible that the stylus on WASP will get blocked.   Over the years we have had a number of questions concerning causes and prevention of stylus blocking.   This technical note identified some of the frequently asked questions and more importantly it provides the answers.

Question 1    Is there any food sample for which WASP is not suitable?

Answer:

Almost all samples can be used in WASP but some will require a large dilution to have a useable viscosity.

NOTE: A possible problem with large dilutions is the samples after dilution may not contain sufficient bacteria for spiral plating therefore care is required to ensure an optimum viscosity/dilution is made in these cases.


Question 2    Is it good to use detergent solution or warm water for washing the stylus tube?

Answer:

For customers using samples that leave a residue in the sample tube we would recommend that the stylus is regularly flushed through with a hot detergent solution i.e. detergent at 50° - 55° and 1% detergent. With WASP set in valve mode draw the detergent solution into the instrument as though it were a normal sample. If you have a large build up it may be necessary to push the solution through the stylus using the “back flush” system and a 50 ml syringe. This requires removing the cover of the instrument, rather than using the ‘power wash’ or ‘sample return’ functions.

Question 3    How do we prepare difficult samples?

Answer:

The steps for preparation of difficult samples are as follows:

- The sample should be homogenised in a Stomacher Filter Bag or blender.

- After stomaching the sample is transferred to a dispo beaker.

- If there are still problems with particulates the sample should be left in the refrigerator for 15 minutes so that the particles can settle to the bottom of the dispo beaker.

- The sample should be drawn into the WASP carefully so that only the liquid part of the sample (the supernatant) is used.

If the sample is still a problem the customer should consider using the syringe mode only to introduce the minimum volume of sample and reduce the risks of blockage around the area where sample leaves the stylus and enters the syringe body and syringe plunger.

What is the largest volume the WASP can dispense?
WASP 400µl DEPOSITION MODE

The basis of the spiral plating technique is that a known volume of sample is dispensed onto a rotating agar plate in an Archimedes spiral. This is accomplished with such precision that the amount of sample on every segment of the plate is known, constant and repeatable. The volume of sample deposited per unit area of plate decreases across the spiral, resulting in a dilution effect. This technique allows for the counting of isolated colonies in specific sectors of the plate where a known volume of sample has been deposited.

All WASP 2 models incorporate a 500µl backfill syringe and different software, making the WASP 2 unique. WASP 2 has many potential deposition volumes. The normal WASP 2 deposition volume options are 50, 100, 200 or 400µl. The product software optimises the speed of turntable rotation according to the volume of sample being dispensed. This is necessary to compensate for centrifugal force, which, with larger volumes, would destroy any gradient in the deposited sample volume. For all but the 400µl option, WASP 2 operates in either linear or logarithmic deposition mode.
 
The amount of sample deposited when the 400µl deposition mode is selected precludes applying a continually reducing gradient. For this reason the 400µl deposition is applied in a linear form only. The available count range is from 7.5 x 102/ml (assuming that 300 colonies on the plate is the maximum number that can actually be counted and distinguished manually) down to 8/ml at the lower end.

Whilst this lower limit is an improvement over the lower limit provided when a 200µl deposition mode is selected, the other benefits unique to the 400µl linear deposition mode are:

- A perfectly even and reproducible lawn of bacteria.
- A reduction in the areas of ‘clumping’ confluent growth that can occur when using a “hockey stick” to spread the sample.
- No growth in the meniscus area of the plate, which enables automatic colony counters to provide more accurate and reproducible results.

In conclusion, the WASP with this unique 400µl deposition mode offers significant advantages over a standard pour or spread plate.