Procedures for the Storage of Microorganisms

Long- and short-term preservation of microorganisms for future study has a long tradition in

microbiology. Culture collections of microorganisms are valuable resources for scientific

research in microbial diversity and evolution, patient care management, epidemiological

investigations, and educational purposes. Preserved individual strains of microorganisms

serve as permanent records of microorganisms’ unique phenotypic profiles and provide the

material for further genotypic characterizations. Such reference collections can encompass

rare infectious agents unique to an individual or catalog the history of disease caused by

common pathogens such as those responsible for community outbreaks.

There are multiple methods for microbial preservation. Effective storage is defined by the

ability to maintain an organism in a viable state free of contamination and without changes

in its genotypic or phenotypic characteristics. Secondly, the organism must be easily

restored to its condition prior to preservation. Microbial preservation methods have been

evaluated extensively over the past 60 years, and often, optimal methods for preservation

depend on a microorganism’s taxonomic classification. Review articles, monographs, and

books have been published that provide detailed information about the storage of various

types of microorganisms (1, 15, 19, 20, 36). For clinical microbiology laboratories, simple

and broadly applied methods are necessary to maintain organisms for short- and long-term

recovery. This chapter presents methods that can be used for the storage of bacteria,

protozoa, fungi, and viruses.

OVERVIEW OF PRESERVATION METHODS Back to top

Short-Term Preservation Methods

Direct Transfer to Subculture

The simplest method for maintaining the short-term viability of microorganisms, most often

used for bacteria, is periodic subculture to fresh medium. Although simple, if microorganisms

are saved for more than 1 week, this method is potentially labor-intensive, requires

extensive laboratory space, and may compromise a microorganism’s phenotypic profile. Each

transfer to a new subculture increases the likelihood of mutation with undesirable changes in

a microorganism’s characteristics.

The interval between transfers varies among organisms. Additionally, the rate of mutation is

quite variable. Some organisms appear stable indefinitely with repeated transfer, and others

may change phenotypic traits after as few as two or three passages. The actual rate of

mutation, however, has not been studied using sequencing technology. Issues that must be

addressed with direct transfer include the medium to be used, the storage conditions, and

the frequency of transfer.

Maintenance Medium

The medium should support the survival of the microorganism but minimize its metabolic

processes and slow its rate of growth. Extreme environments should be avoided because

microorganisms have the unique ability to adapt through mutation events in order to survive

in suboptimal surroundings. A medium with too high a nutrient content will induce rapid

replication that requires more frequent transfers. The optimal medium for maintaining

microorganisms has not been clearly defined and most likely varies from one genus to

another. Media that have been used include distilled water, tryptic soy broth, and nutrient

broths (e.g., from Becton Dickinson and Co. and Oxoid Ltd.), all of which may be used with

or without cryopreservatives.

Storage Conditions

Many laboratories store organisms, most often bacteria, for short periods on routine agar

media at the workbench. Cultures kept in this fashion are subject to drying. A better method

is to transfer organisms into screw-top test tubes and to store them in an organized location

away from light and significant temperature changes. To prevent drying, caps can include

rubber liners, or film can be wrapped over the top of the tube before or after the cap is

screwed on. Storage at lower temperatures (5 to 8°C) slows metabolic processes and

maintains viability for longer periods.

Frequency of Transfer

There is no set protocol for the frequency of transfer since storage conditions, media used,

and types of microorganisms vary among laboratories. Individual laboratories should conduct

studies for each category of microorganism to determine acceptable intervals between

transfers under their conditions used for storage. Such studies would involve performing

subcultures at scheduled times until the laboratory identifies an acceptable interval between

transfers at which a microorganism can reliably and reproducibly be recovered. (When

transfers are performed, 5 to 10 representative colonies should be used to avoid the

possibility of introducing an altered genotypic or phenotypic characteristic.)

Quality Control Procedures

Although it is not necessary with each transfer, the status of the specimen should be

assessed periodically. Ongoing viability, stability of phenotype, microorganism identity, and

the rate of contamination of specimens should be determined and noted in a log.

Immersion in Oil

An alternative to capping tubes is to add a layer of mineral oil to the top of the specimen.

Many bacteria and fungi can be stored for periods of up to 2 to 3 years by this method, and

transfers are not needed as frequently. Microorganisms are still metabolically active in this

environment, and mutations can still occur. Contamination of the specimen can occur if the

mineral oil is not adequately sterilized.

Mineral oil should be medicinal-grade oil with a specific gravity of 0.865 to 0.890 (e.g., from

Roxane Laboratories or Becton Dickinson and Co.). For sterilization, it should be heated to

170°C for 1 to 2 h in an oven (15). Autoclaving is not considered acceptable.

To prepare the specimen, an inoculum of 5 to 10 colonies of the microorganism should be

placed on an agar slant or in tubed broth media. Once growth is identified, a layer of mineral

oil at least 1 to 2 cm deep is added, and the agar must not be exposed to air. As with the

simple transfer method, tests for viability should be performed to determine the optimal

transfer schedule that will ensure microorganism recovery. Transfers will be less frequent

than those of microorganisms stored without oil; however, oil is more difficult to add to vials

and to clean up in the event of spills.

Freezing at −20°C

Refrigeration or freezing in ordinary freezers at −20°C may be used to preserve

microorganisms for periods longer than those that can be accomplished by repeated

transfers. Viability may be maintained for as long as 1 to 2 years for specific

microorganisms, but overall, damage caused by ice crystal formation (20) and electrolyte

fluctuations (15) results in poor long-term survival. The medium used for storage appears to

be important, since preservation times vary from a few months to 2 years depending upon

which medium is used (17, 20, 22). Modern self-defrosting freezers with freeze-thaw cycles

must be avoided because cyclic temperature fluctuation will destroy the microorganism.

Drying

Although most microorganisms do not survive drying, molds and some spore-forming

bacteria may be dried and stored for prolonged periods. Soil can be used as a storage

medium if it is autoclaved and air dried. Soil should be autoclaved for several hours on two

successive days. It is then transferred into sterile glass tubes. A 1-ml suspension of the

microorganism is inoculated into the tube, and the tube is left open to air dry before being

closed with a sterile stopper. The sample is stored in a refrigerator (15). Although potentially

effective, soil is not a standardized, defined, and consistent product for use over long

periods. Instead, commercial silica gel can be used in small cotton-plugged tubes after being

heated in an oven to 175°C for 1.5 to 2 h (20), with moderately successful recovery of fungi.

Alternatively, a suspension of 108 microorganisms can be inoculated onto sterile filter paper

strips or disks. The paper is dried in air or under a vacuum and is placed in sterile vials.

These vials can be stored in the refrigerator for up to 4 years, and then single strips or disks

can be removed as needed (15). This method is commonly used for quality control

organisms.

Storage in Distilled Water

Most organisms do poorly in distilled water, but some survive for prolonged periods. Many

fungi andPseudomonas spp. survive for several years in distilled water at room temperature

(20, 27). McGinnis et al. found that with the exception of fungi that do not easily sporulate,

93% of yeasts, molds, and aerobic actinomycetes can be easily and inexpensively preserved

this way (26).

Long-Term Preservation Methods

Whereas the methods described above may be used to store microorganisms for periods of

up to a few years, ultralow-temperature freezing and freeze-drying (lyophilization) are

recommended for long-term storage. Although the initial investment in ultralow-temperature

freezers and lyophilization may be costly, these methods are less labor-intensive over time,

require less laboratory space (e.g., a cryovial versus broth or agar media), and reduce the

chances of mutation events. Of course, mutations may still occur, and this phenomenon was

observed in Staphylococcus aureus strains that lost the mecA gene during long-term

preservation at −80°C (39). Similar to those with other preservation methods, survival rates

after freeze-drying vary with species. Evaluating microorganisms over a 10-year period,

Miyamoto-Shinohara et al. found that survival rates after freeze-drying

for Brevibacterium spp. and Corynebacterium spp. approached 80%, whereas those

for Streptococcus mutans decreased to 20% after 10 years (27).

Ultralow-Temperature Freezing

Microorganisms can be maintained at temperatures of −70°C or lower for prolonged periods.

Systems for achieving these temperatures include ultralow-temperature electric freezers and

liquid nitrogen storage units. With either system, unwanted heating can occur due to the loss

of electrical power or liquid nitrogen. Close observation of the system and an adequate alarm

mechanism are essential, since any increase in temperature will reduce viability. In the event

that the temperature does rise, restoring power and returning to the target storage

temperature as quickly as possible are essential. The presence of a cryopreservative such as

glycerol may reduce the risk to microorganisms upon short exposure to higher temperatures

(29). If thawing does occur, there are no guidelines for rapid restoration of the storage

condition. Refreezing of the sealed vials as described below may be considered.

Storage Vials

Storage vials must be able to withstand very low temperatures and maintain a seal for their

contents. Plastic (polypropylene) or glass (borosilicate) tubes may be used. Plastic vials with

screw tops and silicone washers are much easier to use than glass vials that must be sealed

with a flame and then scored and broken open. Several commercial suppliers stock

acceptable vials (e.g., Fisher Scientific Products, VWR Scientific, Wheaton Science Products,

and Becton Dickinson and Co.). Vials come in a variety of sizes. Half-dram vials are available

from several suppliers and can be conveniently packaged in a 12-by-12 grid so that 144 vials

are stored in one box or layer.

Cryoprotective Agents

To protect microorganisms from damage during the freezing process, during storage, and

during thawing, cryoprotective agents are often added to the culture suspension. Whereas

most bacteria, fungi, and viruses survive better with such additives, studies have shown that

cryoprotective agents significantly damage others. The reader is referred to detailed

references for specifics (Table 1) (1, 20). Rapid freezing without additives may still be

acceptable for the long-term survival of protozoa, although freeze-drying may be preferred.

Preparation of Microorganisms for Freezing

Microorganisms are inoculated into a medium that adequately supports maximal growth.

Cultures are allowed to mature to the late growth or stationary phase before being

harvested. Broth specimens are centrifuged to create a pellet of microorganisms. The pellet

is withdrawn and resuspended in 2 to 5 ml of broth with the appropriate concentration of

cryoprotectant additive. For agar specimens, broth containing the cryoprotectant is placed on

the surface of the agar. The surface is scraped with a pipette or sterile loop to suspend

microorganisms, and then the broth mixture is pipetted directly into freezer vials.

Alternatively, the agar surface can be scraped with a sterile loop. The microorganisms can

then be transferred directly into the vial of cryoprotectant and emulsified into a final dense

suspension. The volume of the aliquots to be frozen is typically 0.2 to 0.5 ml.

Freezing Method

The American Type Culture Collection (ATCC) recommends slow, controlled-rate freezing at a

rate of 1°C per min until the vials cool to a temperature of at least −30°C, followed by more

rapid cooling until the final storage temperature is achieved (1). Controlled-rate freezers are

required for the initial phase of cooling. Studies in the 1970s showed that uncontrolled-rate

freezing may be acceptable for most organisms and is much less expensive or laborintensive

(20). When organisms are stored in liquid nitrogen, however, it is still

recommended that vials be placed initially in a −60°C freezer for 1 h and then transferred

into the liquid nitrogen. When organisms are stored permanently at −60 to −70°C, the vials

can be placed directly into the freezer.

Small glass beads or plastic beads (e.g., from Fisher Scientific Products or Wheaton Science

Products) can also be added to storage vials before freezing. The culture suspension will coat

the beads, and then individual beads can be removed from storage for reconstitution without

thawing the entire sample (13).

Thawing

Damage to microorganisms occurs as they are warmed from the frozen state. Critical

temperatures appear to be between −40 and −5°C. Studies suggest that rapid warming

through these temperatures improves recovery rates. Stored culture vials should be warmed

rapidly in a 35°C water bath until all ice has disappeared (1, 20). Once a vial is thawed, it

should be opened and the organism should be transferred to an appropriate growth medium

immediately. Great care must be exercised during the thawing phase, since rapid

temperature changes and resulting air pressure changes inside vials can cause the vials to

explode. Protective clothing and eyewear must be worn during this process.

Freeze-Drying (Lyophilization)

Freeze-drying is considered to be the most effective way to provide long-term storage of

most bacteria, yeasts, sporulating molds, and viruses. Better preservation occurs with

freeze-drying than with other methods because freeze-drying reduces the risk of intracellular

ice crystallization, which compromises viability. Removal of water from the specimen

effectively prevents this damage. Among bacteria, the relative viability with lyophilization is

greatest with gram-positive bacteria (sporeformers in particular) and decreases with gramnegative

bacteria (20, 28), but overall, the viability of bacteria can be maintained for as long

as 30 years. In addition, large numbers of vials of dried microorganisms can be stored with

limited space, and organisms can be easily transported long distances at room temperature.

The process combines freezing and dehydration. Organisms are initially frozen and then dried

by lowering the atmospheric pressure with a vacuum apparatus. Freeze-drying has been

extensively reviewed in the past (19), and the required equipment includes a vacuum pump

connected in line to a condenser and to the specimens. Specimens can be connected

individually to the condenser (manifold method) or can be placed in a chamber where they

are dehydrated in one larger air space (chamber or batch method). Alexander et al. and

Heckly have both published detailed descriptions of equipment options (1, 19).

Storage Vials

Glass vials are used for all freeze-dried specimens. When freeze-drying is performed in a

chamber, double glass vials are used. In the chamber method, an outer soft-glass vial is

added for protection and preservation of the dehydrated specimen. Silica gel granules are

placed in the bottom of the outer vial before the inner vial is inserted and cushioned with

cotton. For the manifold method, a single glass vial is used. For both methods, the vial

containing the actual specimen is lightly plugged with absorbent cotton. The storage vial in

the manifold method or the outer vial in the chamber method must be sealed to maintain the

vacuum and the dry atmospheric condition. All vials are sterilized prior to use by heating in a

hot-air oven.

Cryoprotective Agents

Research concerning cryoprotective agents has been extensively reviewed (19). In general,

the two most commonly used agents are skim milk and sucrose. Skim milk is used most

often for chamber lyophilization, and sucrose is used most often for manifold lyophilization.

Skim milk is prepared by making a 20% (vol/vol) solution of skim milk in distilled water. The

solution is divided into 5-ml aliquots and autoclaved at 116°C, with care taken to prevent

overheating and caramelization of the solution. The preparation is then used in smaller

volumes as described above for freezing. Sucrose is prepared in an initial mixture of 24%

(vol/vol) sucrose in water and added in equal volumes to the microorganism suspension in

growth medium to make a final concentration of 12% (vol/vol).

Preparation of Microorganisms for Lyophilization

As with simple freezing, maximum recovery of organisms is achieved by using

microorganisms in the late growth or stationary phase from the growth of an inoculum in an

appropriate growth medium. High concentrations of microorganisms are considered to be

important. The ATCC recommends a concentration of at least 108 CFU/ml (1), and Heckly

suggests a concentration of 1010 CFU/ml or higher (20).

Freeze-Drying Methods

In the chamber method, inner vials with the microorganism suspension are placed in a single

layer inside a stainless steel container. This container is placed in a low-temperature freezer

at −60°C for 1 h. The container is then transferred to a chamber containing dry ice and ethyl

Cellosolve (Becton Dickinson and Co.) and covered with a sealable vacuum top, which is

connected in sequence to a condenser reservoir also filled with dry ice and ethyl Cellosolve

and to a vacuum pump. The vacuum is maintained at a minimum of 30 μm Hg for 18 h. At

the same time, the outer vials are prepared by being heated in an oven overnight, filled with

silica gel granules and cotton, and placed in a dry cabinet (with <10% relative humidity).

The freeze-dried inner vials are inserted into the outer vials, and the outer vials are heat

sealed. Multiple different strains or species should probably not be processed in the same

batch. Cross contamination rates vary from 0.8 to 3.3% when two different microorganisms

are placed on opposite sides of the same container and are as high as 8.3 to 13.3% when

microorganisms are intermingled (3).

In the manifold method, a rack of individual vials is used rather than a single container. The

rack is placed in a dry ice-ethyl Cellosolve bath. After the freezing process, the vials are

connected by individual rubber tubes in sequence to the condenser container filled with dry

ice and ethyl Cellosolve and to the vacuum pump. As in the method described above, the

vacuum is maintained at 30 μm Hg for 18 h and then the individual vials are sealed.

Storage

Individual vials need to be appropriately labeled and sorted. Storage at room temperature

does not maintain viability and is not recommended. Storage at 4°C in an ordinary

refrigerator is acceptable, but survival rates may be improved at temperatures of −30 to

−60°C (1, 19).

Reconstitution

Care must be taken when opening vials for reconstitution because of the vacuum inside the

vial. Safety glasses should always be worn, and vials should be covered with gauze to

prevent injury if the vial explodes when air rushes in. Reconstitution should also be

conducted in a closed hood to avoid dispersal of microorganisms. The surface of the vial

should be wiped with 70% alcohol, and then the top of the glass vial can be scored and

broken off or punctured with a hot needle. A small amount (0.1 to 0.4 ml) of growth medium

is injected into the vial with a needle and syringe or a Pasteur pipette, the contents are

stirred until the specimen is dissolved, and then the entire contents are transferred with the

same syringe or a pipette to appropriate broth or agar media. A purity check must be done

on each specimen because of the possibility of either cross contamination or mutation during

the preservation process.

Newer Technologies

The long-term preservation methods previously described are specifically designed for

recovery of microorganisms for further cultivation. Culture-independent tests based on

antigen or nucleic acid technologies are in widespread use and do not require viable

microorganisms. In this regard, storage of microorganisms to preserve their antigens or

nucleic acids is also important for clinical laboratories. The use of Whatman Flinders

Technology Associates (FTA) matrix cards (Whatman International Ltd., Maidstone, United

Kingdom) or other filter paper-based products is a novel approach for long-term storage of

microbial DNA that is safe (microorganisms are inactivated), inexpensive, and fast (4, 33).

Bacterial and/or fungal cell suspensions are applied directly to dry FTA paper. The FTA cards

are impregnated with buffers, free radical trap and protein denaturants that lyse cell

membranes on contact, entrap DNA, and protect DNA from degradation. This technology has

been successfully applied to a variety of bacteria and fungi and serves as a reusable DNA

archiving system. Although beyond the scope of this chapter, direct specimens such as blood

can be preserved using a dry blood spot on filter paper or with a non-paper-based matrix for

future antibody or nucleic acid testing to detect human immunodeficiency virus (5, 8, 25),

hepatitis B virus (8), hepatitis C virus (8, 25), Rickettsia typhi, and Orientia

tsutsugamushi (30).

Procedures for Specific Organisms

Procedures for specific organisms are described below and summarized in Table 1.

Bacteria

All of the material presented in this chapter applies primarily to the preservation of bacteria.

Simple transfer, storage under mineral oil, drying, or freezing at −20°C can maintain

bacteria for short periods; freezing in ultralow-temperature electric freezers at −70°C or in

liquid nitrogen at −196°C or freeze-drying can provide long-term preservation. A summary

of the studies of bacterial preservation has been published (20). In general, serial transfer

will preserve bacteria for up to a few months, storage under mineral oil or with drying will

last 1 to 2 years, freezing at −20°C will preserve bacteria for 1 to 3 years, freezing at −70°C

will preserve bacteria for 1 to 10 years, and freezing in liquid nitrogen and freeze-drying will

preserve bacteria for up to 30 years (15). For fastidious bacteria such as Streptococcus

pneumoniae, Neisseria spp., and Haemophilus spp., the optimal methods are lyophilization

and freezing at −70°C by using Trypticase soy broth with glycerol as a preservation medium

(31, 35, 40). Stock cultures of quality control microorganisms can be maintained in a

cryopreservative suspension for up to 1 year at −20°C or indefinitely at −70°C.

Protozoa

Information concerning the preservation of protozoa is limited, in keeping with the infrequent

need for such a process in clinical microbiology laboratories. Variable methods for individual

genera are described. In general, freezing appears to be preferred to freeze-drying. All of the

following procedures are as described by the ATCC (1).

Acanthamoeba spp., Leishmania spp., Naegleria spp., Trichomonas spp.,

and Trypanosoma spp. can be handled as described above for ultralow-temperature freezing

with 5% (vol/vol) DMSO as the cryoprotective agent. These organisms should be stored in

liquid nitrogen.

Acanthamoeba spp. and Naegleria spp. can also be dried at room temperature onto filter

paper. Aliquots of a microorganism suspension (0.3 ml) are pipetted onto the paper in a shell

vial and dried in air for 14 days at room temperature and then in a vacuum desiccator for an

additional week. The vials are sealed and stored in liquid nitrogen.

Entamoeba spp. are stored frozen at −40°C. Specimens should be suspended in a mixture of

growth medium containing 12% (vol/vol) DMSO and 6% (vol/vol) sucrose.

Leishmania spp. may also be prepared by inoculation of the organism into an animal host. At

the peak of infection, the spleen is harvested and homogenized in half the final volume of

ATCC medium 811 salt solution. Freezing is completed with 10% glycerol as the

cryoprotectant.

Plasmodium spp. can be stored from infected blood samples. At the height of parasitemia,

blood is obtained and anticoagulated with the following preparation: 1.33 g of sodium citrate,

0.47 g of citric acid, 3.00 g of dextrose, 200 mg of heparin (sodium), and 100 ml of distilled

water. The final concentration of anticoagulant added to blood is 10%. To this anticoagulated

blood, 30% glycerol in 0.0667 M phosphate buffer is added to a final concentration of 10%

(vol/vol) glycerol. Freezing should occur in liquid nitrogen.

Trypanosoma spp. must be harvested from an animal host. At the peak of parasitemia, blood

is withdrawn into heparinized tubes and diluted 1:1 in Tyrode’s solution (8.0 g of NaCl per

liter, 0.02 g of KCl per liter, 0.2 g of CaCl2 per liter, 0.1 g of MgCl2 per liter, 0.05 g of

NaH2PO4 per liter, 1.0 g of NaHCO3 per liter, and 1.0 g of glucose per liter) with 1 to 5%

phenol red added. Then 5% DMSO is added as the cryoprotectant, and the specimen is

stored in liquid nitrogen.

Yeasts and Filamentous Fungi

All of the techniques described above have been applied to the storage of yeasts and fungi

(7, 15, 20, 36). The individual method employed depends upon the species to be preserved

and whether or not it sporulates.

Subculturing. Subculturing is the simplest method of maintaining living fungi and involves

serial transfer to fresh solid or liquid media. Storage is accomplished usually at room or

refrigerator temperature. Fungi may be maintained by subculturing for a number of years.

Care must be taken to avoid aerosolization and contamination of the laboratory or other

specimens.

Storage under oil. Whereas species of Aspergillus and Penicillium have remained viable

under oil for 40 years (36), many species have shown deterioration after 1 to 2 years and

must be transferred periodically. Taddei et al. also reported the successful storage and

recovery of actinomycetes stored under paraffin oil for 10 to 30 years (37).

Water storage. Many fungi can be stored successfully for prolonged periods in distilled

water (27, 32). A simple method is to pipette 6 to 7 ml of sterile distilled water onto 2-weekold

culture slants in screw-cap tubes. The spores and fragments of hyphae are dislodged by

scraping with the pipette, and the suspension is transferred to a sterile 1-g vial, which is

tightly capped and stored at 25°C. Fungi are revived by subculturing 0.2 to 0.3 ml of the

suspension to appropriate media (6).

An alternative method is to cut agar blocks from the growing edge of a fungal colony and

place them in sterile distilled water in bottles with screw-cap lids (18). The cultures are

stored at 20 to 25°C. The fungi are retrieved by removing a block and placing it mycelium

side down on growth medium appropriate for that species (36). Contamination (22.8%) is a

significant problem with this method (18).

Drying. Drying as described above has been used for fungi. Only 6 of 16 genera of fungi

stored in this fashion survived for 4 years (2). The greatest success is reported for

sporulating fungi stored in silica gel or in soil (36).

Freezing. Fungi have been successfully preserved by storage in liquid nitrogen by using

glycerol or DMSO as a cryopreservative. Broth cultures containing nonpathogenic fungi are

disrupted in a Waring blender and suspended in equal parts of DMSO or glycerol to achieve

final concentrations of 5 or 10%, respectively. Pathogens should not be disrupted in a

mechanical blender because of the potential biohazard associated with

aerosolization. Histoplasma, Paracoccidioides, and Blastomyces species should be frozen in

the yeast phase, and Coccidioides species should be frozen in the early mycelial phase to

minimize exposure of laboratory personnel. Otherwise, procedures for freezing are as

described above.

Freeze-drying. Most spore-forming fungi can be preserved by freeze-drying. Cultures to be

stored by freeze- drying should be grown on agar or broth media to the point of maximum

sporulation (1) and processed as described above. Survival in storage for many years has

been demonstrated (11, 34), but this is true only for sporulating organisms. Young

vegetative hyphae of fungi do not survive freeze-drying (36).

Viruses

Viruses tend to be more stable than other microorganisms because of their small size and

simple structure and the absence of free water. Many viruses can be stored for months at

refrigerator temperatures or for years by ultralow-temperature freezing or freeze-drying.

Storage at −20°C is not recommended (20, 23). Larger viruses tend to be less stable than

smaller ones (16).

Ultralow-temperature freezing is effective in a number of situations. In addition to

cryoprotectants described above, sucrose-phosphate-glutamate containing 1% bovine

albumin (SPGA) (20, 23) and hypertonic sucrose are particularly effective, the latter for

storing labile viruses such as respiratory syncytial virus (24). If ultralow-temperature

freezing is employed, the rate of freezing should be as high as possible, using small-volume

suspensions (0.1 to 0.5 ml). In addition to freezing of pure isolates, stool specimens known

to contain viral enteric pathogens have been maintained at −70 to −85°C for 6 to 10 years

with reasonable recovery and no change in the morphological characteristics of astroviruses,

small round structured viruses, enteric adenoviruses, rotaviruses, and caliciviruses (41).

Gallo et al. evaluated five types of media for storage of human immunodeficiency virusinfected

peripheral blood lymphocytes and concluded that freezing peripheral blood

lymphocytes in RPMI 1640 containing 10% fetal bovine serum and 10% DMSO and storing

them at −60°C is acceptable for human immunodeficiency virus isolation (14).

Freeze-drying is probably the optimum method for preserving viruses for extended periods.

A detailed review of acceptable procedures has been published (16). Virus suspensions

freeze-dried in medium supported with SPGA appear to survive better (20, 38).

Lyophilization of polioviruses and other enteroviruses works best when electrolytes are

removed by dialysis or ultrafiltration (20).

Select Agents

In response to the Public Health Security and Bioterrorism Preparedness and Response Act of

2002, federal regulations require laboratories that store select agents to register and comply

with the standards established by the act (12). A current and complete list of

microorganisms considered to be select agents can be found

at www.cdc.gov/od/sap. Regardless of the method for long-term preservation, laboratories

must register with the Department of Health and Human Services and Centers for Disease

Control and Prevention Select Agent Program. In order to minimize risk to public health and

safety, select agents must be stored in a highly secured area with restricted access and

appropriate safeguards. Only registered individuals who have completed training for handling

select agents can access and retrieve these microorganisms from storage. An accurate and

current inventory of select agents held in long-term storage must be maintained.