Which kind of lab is supposed to handle bacteriophages?

Bacteriophages are viruses that affect bacteria and not plants, human or any other mammals, Due to their specificity, there fore it pose no harm to other creatures except that specific bacteria host. To determine whether which laboratory can handle which bacteriophage it depends on the host used instead of the phage handled, there fore the determinant is bacteria and not the virus. For example bacteriophages for Mycobacterium must be handled in higher labs due to the fact that at one point the host (which is the deadly bacteria) will be used.  Lets polish our knowledge on bio-safety laboratory levels.

 Bio-safety Level 1

Biosafety level one, the lowest level, applies to work with agents that usually pose a minimal potential threat to laboratory workers and the environment and do not consistently cause disease in healthy adults. Research with these agents is generally performed on standard open laboratory benches without the use of special containment equipment. BSL 1 labs are not usually isolated from the general building. Training on the specific procedures is given to the lab personnel, who are supervised by a trained microbiologist or scientist.

Standard microbiology practices are usually enough to protect laboratory workers and other employees in the building. These include mechanical pipetting only (no mouth pipetting allowed), safe sharps handling, avoidance of splashes or aerosols, and decontamination of all work surfaces when work is complete, e.g., daily. Decontamination of spills is done immediately, and all potentially infectious materials are decontaminated prior to disposal, generally by autoclaving. Standard microbiological practices also require attention to personal hygiene, i.e., hand washing and a prohibition on eating, drinking or smoking in the lab. Normal laboratory personal protective equipment is generally worn, consisting of eye protection, gloves and a lab coat or gown. Biohazard signs are posted and access to the lab is limited whenever infectious agents are present.

Bio-safety Level 2

Biosafety level two would cover work with agents associated with human disease, in other words, pathogenic or infectious organisms posing a moderate hazard. Examples are the equine encephalitis viruses and HIV when performing routine diagnostic procedures or work with clinical specimens. Therefore, because of their potential to cause human disease, great care is used to prevent percutaneous injury (needlesticks, cuts and other breaches of the skin), ingestion and mucous membrane exposures in addition to the standard microbiological practices of BSL 1. Contaminated sharps are handled with extreme caution. Use of disposable syringe-needle units and appropriate puncture-resistant sharps containers is mandatory. Direct handling of broken glassware is prohibited, and decontamination of all sharps prior to disposal is standard practice. The laboratory’s written biosafety manual details any needed immunizations (e.g., hepatitis B vaccine or TB skin testing) and whether serum banking is required for at-risk lab personnel. Access to the lab is more controlled than for BSL 1 facilities. Immunocompromised, immunosuppressed and other persons with increased risk for infection may be denied admittance at the discretion of the laboratory director.

BSL 2 labs must also provide the next level of barriers, i.e., specialty safety equipment and facilities. Preferably, this is a Class II biosafety cabinet or equivalent containment device for work with agents and an autoclave or other suitable method for decontamination within the lab. A readily available eyewash station is needed. Selfclosing lockable doors and biohazard warning signs are also required at all access points.

Bio-safety Level 3

Yellow fever, St. Louis encephalitis and West Nile virus are examples of agents requiring biosafety level 3 practices and containment. Work with these agents is strictly controlled and must be registered with all appropriate government agencies.2 These are indigenous or exotic agents that may cause serious or lethal disease via aerosol transmission, i.e., simple inhalation of particles or droplets. The pathogenicity and communicability of these agents dictates the next level of protective procedures and barriers. Add to all the BSL 2 practices and equipment even more stringent access control and decontamination of all wastes, including lab clothing before laundering, within the lab facility. Baseline serum samples are collected from all lab and other at-risk personnel as appropriate.

More protective primary barriers are used in BSL 3 laboratories, including solid-front wraparound gowns, scrub suits or coveralls made of materials such as Tyvek® and respirators as necessary. Facility design should incorporate self-closing double-door access separated from general building corridors. The ventilation must provide ducted, directional airflow by drawing air into the lab from clean areas and with no recirculation.

Bio-safety Level 4

Agents requiring BSL 4 facilities and practices are extremely dangerous and pose a high risk of life-threatening disease. Examples are the Ebola virus, the Lassa virus, and any agent with unknown risks of pathogenicity and transmission (Laboratory insider listed top ten of the most lethal viruses here). These facilities provide the maximum protection and containment. To the BSL 3 practices, we add requirements for complete clothing change before entry, a shower on exit and decontamination of all materials prior to leaving the facility.

The BSL 4 laboratory should contain a Class III biological safety cabinet but may use a Class I or II BSC in combination with a positive-pressure, air-supplied full-body suit. Usually, BSL 4 laboratories are in separate buildings or a totally isolated zone with dedicated supply and exhaust ventilation. Exhaust streams are filtered through high-efficiency particulate air (HEPA) filters, depending on the agents used.

Biosaftety lab levels infographic by CDC
Biosafety lab levels by CDC


Can genomics/meta-genomics replace transimission electron microscopy (TEM)?

Can genomics replace electron microscopy?

next-generation sequencing
next-generation sequencing
This might be suggested by the rise of rapid sequencing and the ensuing increased availability of completely sequenced virus genomes. It is indeed advocated in discussions by unconditional partisans of genomics. The answer is round ‘‘no.’’ Genomics gives us the genome and genes, thus the elementary building blocks of a virus. It also gives gene order and direction of transcription, and it identifies genes coding for proteins with homology to known enzymes or virion components, restriction-modification enzymes, capsid protein size, or the length of tape measure proteins. Further, genomics indicates horizontal gene transfer or gene swapping, may indicate relationships between virus groups and individual viruses and allows for quantification of relationships and the construction of phylogenetic trees. All this provides unprecedented insights into virus evolution and is a precious help in phage classification.
However, electron microscopy provides information on virion structure, while genomics does not show the whole virus, gives not a single dimension, provides no information on virus structure and physicochemical properties, does not identify unusual bases such as 5-hydroxymethylcytosine, and predicts only some biological properties, such as a lysogenic nature. No sequence can indicate simple things such as the size of phage capsids, their geometry, or the number of capsomers. If, as likely, the length of phage tails depends on the length of ruler protein genes (Katsura and Hendrix, 1984; Pedulla et al., 2003), this must be ascertained by the measurement of many phage tails under strict magnification control. Unfortunately, this has not been the case. If, as pretended, a genome contains all information on a virus, we have not yet found the instruction manual to read it. 
With respect to virus identification, genomics generally does not indicate to which virus family a tailed phage belongs; for example, there are no sequences specific to Myo-, Sipho-, or Podoviridae. Only in the case of small polyhedral or filamentous phages (Micro-, Levi, and Inoviridae) does genomics allow for the identification of virus families (Ackermann and Kropinski, 2007). Similarly, a Bacillus tectivirus from the earthworm gut was identified by genomics alone without the benefit of electron microscopy (Schuch et al., 2010). However, in a general way, investigation of a complete virus sequence may take months and is infinitely slower and more labor-intensive than electron microscopy. 

Can metagenomics replace electron microscopy?

 The answer is ‘‘no’’ again. For virus identification, metagenomics relies totally on known and identified genes and genomes, which, in turn, belong to viruses known and characterized by electron microscopy. In other terms, the vast majority of countless genes detected by metagenomics can be identified only to the extent as they belong to known sequences from known viruses. Further, metagenomics will not tell whether any detected sequences belong to complete, infectious virions or not. 

Can electron microscopy replace genomics?

 The answer is ‘‘yes,’’ but only when it comes to the identification of high-level taxonomic categories. Clearly, electron microscopy and genomics (or metagenomics) are not alternatives, but complementary. Both of them answer different questions and appear as different fingers of the same hand.
(Read more about bacteriophage here)

Problems of electron microscopy (TEM) for bacteriophages

Transmission electron microscopy history

Transmission electron microscope
Transmission electron microscope
Electron microscopy always had problems of imaging and interpretation, but the rise of digital electron microscopy and CCD cameras in the 1990s created a novel situation. In a general way, it appears that the quality of phage electron microscopy (read more about phages here) has slipped and that many present-day phage electron micrographs are far inferior in quality to the first images of negatively stained phages taken in the late 1950s (Brenner et al., 1959). A peak in phage electron microscopy was reached in the 1970s (see Dalton and Haguenau, 1973), but this seems to be forgotten. For example, in a personal survey of about 130 phage papers since 2006, which described novel phages by mostly digital TEM, 70 featured low-contrast, unsharp, astigmatic, poor to very poor pictures. Some ‘‘phage descriptions’’ reported neither phage dimensions nor stains and did not even specify the electron microscopes used. Only some 20 papers showed good-quality figures. The decline of phage electron microscopy may be linked to personal factors, namely the loss of great electron microscopists such as Eduard Kellenberger or Tom Anderson, their replacement by inexperienced investigators, and perceived leniency even of reputed journals to accept substandard micrographs. Indeed, regardless of the electron microscope used, poor micrographs can be associated with an inadequate technique, whether in specimen processing or imaging. Digital TEMs and CCD cameras are here to stay. CCD cameras have largely obviated darkroom photography and are wildly popular with inexperienced microscopists who fear work in the darkroom. 
  1. Compared to conventional TEMs, digital electron microscopes appear to be more expensive, cannot be maintained normally by users, and need expensive service contracts. 
  2. Their life span remains to be seen and they are more difficult to control than ‘‘manual’’ TEMs with respect to contrast and magnification. However, conventional photographic chemicals and papers may be difficult to find because the market has shrunk. Bacteriophage Electron Microscopy 23 
  3. The relative quality of the various digital TEMs and CCD cameras is difficult to evaluate in the absence of comparative studies. It seems that present top-grade TEMs, whether produced by FEI, JEOL, or Hitachi and concomitant CCD cameras are roughly equivalent with respect to resolution. The instruments are improved continuously. For example, TEMs manufactured by the FEI Company (Hillsboro, OR), which acquired the Philips Electron Optics Division, produces micrographs of striking quality. 
  4. With ‘‘manual’’ TEMs, contrast is controlled in the darkroom by means of graded filters and papers. In the case of digital TEMs, one can obtain high-resolution and high-contrast pictures by the adjustment of pixel intensities with CCD camera software (Tiekotter and Ackermann, 2009). It is unfortunate that the manufacturers of electron microscopes have seemingly neglected to issue guidelines for contrast enhancement, leaving users to fend for themselves. 
  5. With both ‘‘manual’’ and digital TEMs, magnification is controlled by means of test specimens, for example, catalase crystals (Luftig, 1967) or T4 phage tails. Latex spheres or diffraction grating replicas are suitable for low magnification only (10–30,000x). With ‘‘manual’’ microscopes, magnification can be corrected in the darkroom within minutes. The magnification of digital electron microscopes is normally set by the installer and cannot, or only with great difficulty, be adjusted by the user. To control magnification, the user must photograph test specimens and define correction factors by calculation. 
Practically, it is recommended that 
  1. TEM manufacturers publish instructions for contrast enhancement. 
  2. All specimens are purified before the examination (read about purification here). Crude lysates are to be banished. Purification is achieved most easily by differential centrifugation and washing in the buffer. 
  3. Improve contrast of digital microscopes via Photoshop technology. 
  4. Control magnification regularly by means of test specimens.

In a general way, it appears that the quality of phage electron microscopy has slipped and that many present-day phage electron micrographs are far inferior in quality to the first images of negatively stained phages taken in the late 1950s (Brenner et al., 1959). Credit to Ackerman et al


How to make an outstanding media (agar) plate


Making agar plates, whether they contain TSA, MHA, LB, M9, or any other medium, is a simple procedure. But there are a few finer points that will kill your experiment, make a mess, or just cause you inconvenience if you get them wrong. So let’s put on the record exactly how to make the perfect agar plate for those of you who are
 new to the world of working with bacteria.
media preparation
media preparation

Follow these steps and you’ll get great, or even perfect, agar plates – with no lumps, bubbles, or excess moisture – every time.

Some tips for Pouring Perfect Agar Plates Every Time

1. Use a Recipe

Make up the medium according to the recipe, then add the desired amount of agar (normally about 1% w/v) and stir. If you autoclave without stirring, with the agarose still floating on top of the liquid, you get an agarose cake in the medium. Interesting, but useless.

When making up the agar, only use 3/4 of the volume of the bottle. This allows space for bubbles to rise while the agar is melting in the microwave (and saves you cleaning up overflowing agar from the microwave!).

2. Autoclave

Autoclave your medium for 25 minutes. After autoclaving, you can of course store the medium-agar mix in a toughened glass bottle then melt it in a microwave or water bath when needed. Make sure you use toughened glass bottles, or disaster can strike.

3. Cool It!

Cool the medium-agar mix to 55°C. For routinely consistent results, do the cooling for a couple of hours in a 55°C water bath. Agar starts to solidify at about 50°C. Using the water bath means you can consistently cool the mixture to just above the solidification temperature.

Before I used a water bath, I used to just cool it in the air, but would inevitably forget about it and come back to find solidification had already started – lumpy plates are no good for spreading!

4. Supplement It

You can now add any antibiotics or supplements, and be confident that the agar is at a suitable temperature because you have cooled it in the water bath.

5. Pour the Plates

Use about 30 mL of the agar-medium mix for each plate when using a 100 mm diameter plate. The less agar-medium mix in each plate, the more easily they will dry out. 30 mL is a good amount for long-term storage, 10–20 mL is fine if you are going to use the plates relatively soon.

For consistency, I’d recommend using a serological pipette. Suck up 2–3 mL more than you need to minimize blowing bubbles into the plate.

6. Let It Set

If there are any bubbles in the plates, briefly pass the flame over to pop them. Classic error: trying to move the plates before they’ve set is just asking for trouble. Just leave them alone (and maybe admire your perfect agar plates while you wait)!

7. Get Dry

Dry the plates in the laminar flow hood with the lid slightly off for 30 minutes (or in a 37°C incubator for 2–3 hours, or room temperature for 2–3 days). Drying the plate is very important for storing the plates and growing colonies on them.

If you don’t dry the plates, the moisture will evaporate and condense on the lid during storage or incubation and give you horrible wet plates. At worst the moisture can affect the plating of your cells. Use a timer to remind you when the 30 minutes are up as – in my experience – it is very easy to forget about your plates and come back to find your plates have turned into agar crisps/chips. Tasty.

8. Use It or Store It

Once you’ve poured your perfect agar plates, you can use them immediately or seal them for later use. You can use Parafilm or pop them in the bag that the plates came in for easy storage. Store the plates at 4°C. Guidelines suggest using agar plates within approximately 2 to 4 weeks.

Depending on the additives you have included, the shelf life of the prepared plates might be shorter – make sure you check this before you start so you don’t end up wasting your time (and resources) making too many plates.

A quick way to label your plates is to have a color code for each antibiotic and medium type you tend to use (e.g. red for ampicillin, black for kanamycin, green for LB, blue for M9). Stack the plates and use the appropriately colored lab marker to draw a line down the whole stack. Make sure you keep the color code to hand though.

Now you should have perfect agar plates every time. If you’ve got any further ideas or additions to this protocol, please leave a comment.

Phage work can be done using a lot of different media depending on what is available to the researchers doing the experiment although LB and TSB(A) is the most preferred

Agar Overlay (double layer agar) technique

This technique allows you to produce a homogeneous lawn of bacteria within a thin layer of agar across the surface of a plate. Bacteria are added to a soft top agar (0.75% agar, as opposed to the usual 1.5% for agar plates) which has been melted at 100°C and cooled to 45°C. This is warm enough so the agar remains liquid, but cool enough so that the bacteria are not killed (for a period of time). The melted agar/bacterial suspension is mixed and poured evenly across the top of an agar plate and allowed to solidify.

The bacteria distributed through the top agar will grow to produce a homogeneously turbid lawn. If the freshly seeded lawn is exposed to various antibacterial agents and then incubated at 37°C, any inhibition of bacterial growth will cause a reduction in the turbidity of the lawn near the agent: the greater the antibacterial action, the wider the zone of inhibition. Thus, the antibacterial strength of the agent may be judged by the width of the zone of inhibition around it.

Illustrations of the experimental plan for the prep of the inoculated melted agar, and of the pouring of the inoculated agar onto the pre-warmed plate

EQUIPMENT:
sterile capped 13×100 mm tubes
sterile pipettes (0.1, 1.0 mL) or
sterile tips for Displacement pipetter
hot block, 45°C, warmed up
vortex
Bunsen Burner

SUPPLIES:
melted top agar, about 60°C
fresh overnight culture of indicator bacteria
(such as E. coli B or Staphylococcus aureus)
pre-warmed nutrient agar plates
(or tryptone soy agar plates, etc)
dishpan with diluted Lysol into which to discard used tubes

PROTOCOL:

  1. THE PREVIOUS NIGHT:   Inoculate about 2-3 mL of nutrient broth or tryptone soy broth with the desired indicator bacterium (often E. coli B). Grow as a stationary culture ON at 37°C in the hot block.
  2. THE DAY OF USE:   Pipet about 2 mL of hot melted top agar into sterile capped 13×100 mm tubes in a 45°C hot block. Allow cooling to 45°C (several minutes).
  3. Pipet about 0.1 mL of an ON culture of indicator bacteria into the melted agar, here using an Eppendorf Repeat Pipetter. Down the inside of the tube is OK.
  4. Overlay technique:

    a. Vortex to mix the bacteria into the melted top agar
    b. Immediately pour out onto a pre-warmed agar plate to empty the tube.
    c. Immediately tilt back and forth, shake gently to evenly distribute. Avoid bubbles, and stop agitating before agar begins to gel. Let set undisturbed to gel fully, (several minutes.)
  5. When fully gelled, assign and label positions on the plate bottom where agents will be applied or operations performed such as application of antibacterial agents, antibiotics, exposure to UV, etc. Apply liquids to 5 mm sterile filter discs placed on top of the top agar. Do not let run.
  6. Invert, incubate overnight at 37°C.
  7. THE NEXT DAY:  Next AM, read the plates. Where growth is thickest, there was the least antibacterial action. Where the thinnest, the greatest. Illustrate the plate, measure the zones of inhibition (width of the zone from the edge of agent to the edge of a zone), record data.

The Double-Layer Agar (DLA) technique is extensively used in phage research to enumerate and identify phages and to isolate mutants and new phages.

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