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One of the three domains of life (the others being Eukarya and ARCHAEA), also called Eubacteria. They are unicellular prokaryotic microorganisms which generally possess rigid cell walls, multiply by cell division, and exhibit three principal forms: round or coccal, rodlike or bacillary, and spiral or spirochetal. Bacteria can be classified by their response to OXYGEN: aerobic, anaerobic, or facultatively anaerobic; by the mode by which they obtain their energy: chemotrophy (via chemical reaction) or PHOTOTROPHY (via light reaction); for chemotrophs by their source of chemical energy: CHEMOLITHOTROPHY (from inorganic compounds) or chemoorganotrophy (from organic compounds); and by their source for CARBON; NITROGEN; etc.; HETEROTROPHY (from organic sources) or AUTOTROPHY (from CARBON DIOXIDE). They can also be classified by whether or not they stain (based on the structure of their CELL WALLS) with CRYSTAL VIOLET dye: gram-negative or gram-positive.
It can produce blueish-green pigment in nutrient broth. Can utilize gelatin. Can produce H2S in SIM medium, urease positive, citrate and catalase positive. And it is also a phosphate solubilising bacteria, can produce IAA.
I am surprised that so many suggested elaborate test when you can get for much less effort and money / time the definite answer using 16S rRNA sequence (Cost of sequence is about US $ 5 -29 , depending on the service, If the sequence is between two species you can do --based on Bergey's-- very few differentiating tests to nail it down. Don't forget you also could have a novel taxa and you don't find that out by just doing the suggested test.. Don't think in a box. -
Has you anyone tried MATH test (microbial adhesion to hexadecane)?
Mar 6, 2013
I searched for this method on pubmed but there is no common rule for using this method. Added hexadace amount, reading absorbance and incubation time and temperature are different from each other in different papers. Has anyone ever experienced this test for salmonella strains?
i was read some research paper such as Bio degradation of polyethylene by the thermophilic bacterium Brevibacillus borstelensis
D.Hadad , s. Geresh and A Sivan. journal of Appllied Microbiology 2005 ,98,1093-1100
It is seen that generally people sequence 16S rDNA for the identification of a bacterium. However, it has low phylogenetic power at the species level (Janda and Abbott, 2007) and cannot be used to discriminate species of some genera. How can the identity of a bacterium be resolved to a species or subspecies level?
First, there are no clear (official) rules on species/subspecies determination. So you'll have to go along with the current determination of the species/subspecies in the genus you're interested in.
Second, whether your isolate belongs to a certain species or subspecies depends on the results of different tests and depends on the genus. In Deinococcus, 16S rRNA gene sequencing will satisfy to characterize the isolate at species level. For others you could use MLSA, MLVA or even ANI analysis. These will resolve the isolates of the genus to a certain level, at which point you might have to consider to change the current opinion on species/subspecies classification in that genus.
From my experience, I believe that MLSA or MLVA can do the trick, but again, that depends strongly on the genus of interest. If you have plenty of resources you could do whole genome sequencing and run an ANI analysis. Other tests like PFGE, RAPD, AFLP, RFLP or REP-PCR can also help you but their resolution is generally lower than MLSA or MLVA. MLSA is easier to design but slightly more expensive than MLVA.
I want to calculate the sequence identity of some 16S sequences to see if I've reached the species cut-off of 97% of identity. I tried with BLAST and SIAS (http://imed.med.ucm.es/Tools/sias.html), but the results are different...any suggestions?
Sequence identity values from multiple sequence alignments are more reliable. Clustal is a program for generating multiple sequence alignments . SIAS is simple and good but you can do the same with ClustalX. Do the alignment or load the alignment in ClustalX and then under trees chose output tree format options and select %identity matrix.
I plan to extract an over-expressed protein that has a specific structure from BL21. We have seen it through immunofluorescence microscopy. Sonication cannot be used since it may destroy the structure of the protein. It's difficult to find a proper lysis buffer. I plan to use Lysozyme and Triton X-100 with 1mM PMSF, in GTE buffer (Glucose, Tris-HCl and EDTA), but it's hard for me to determine the concentration and reaction conditions. Could anyone give me some suggestions?
Thanks a lot to all the answers. Maybe I failed to explain something accurately. The structure we are interested in is a complex of the over-expressed protein. We plan to do Cryo-EM after that. Thus I think sonication may destroy its structure.... Just a guess. French Press is a really good idea and we have the machine. I'll have a try.
I've done many protein purifications with both soluble and very insoluble proteins. I have never had a problem with loss of activity or structure using sonication (@ 4C on ice, 3 ½ min total time 4 passes at 30s each with 30s rests in between - slowly move sonicator tip up/down while sonicating without touching the bottom or sides of the tube. Start at level 5 and slowly turn dial up each time until the total time has elapsed) and have found that it is very comparable to results lysing with a french press (@ 4C, 2 passes at 1.5 ml/min). However the method that you choose would depend on your protein because sonication can sometimes result in the formation of aggregates.
As far as your lysis buffer selection goes you can try the following, however this will depend on the purification method you are using and what tag is on your protein (ie. His6, GST etc.): 50mM Tris-HCl, 300mM NaCl **this can be varied from 100-500mM**, 20mM Imidazole ** this can be varied from 20-40mM**, 0.05% β-mercaptoethanol, 0.5% triton X-100 ** this can be varied from 0.4-2%** Set pH8.0, filter sterile with 0.22um. Before use add the following protease inhibitors: 1mM PMSF, 0.5µg/ml Leupeptin, 10 µg/ml Aprotinin (or can use the Roche protease inhibitor tablets as previously mentioned before). Do not use strong reducing agents such as DTT, DTE, EGTA or EDTA with Ni-NTA resin because this can dramatically reduce the metal binding capacity towards your protein or strip the Ni2+ from the column. In addition, Its is important to properly prepare your beads prior to avoid any protein degradation. If you are planning on performing activity assays...i would avoid the use of phosphate buffers.
Is your protein easy or difficult to solubilize? There are a few suggestions i could give if your protein is mostly found in the pellet after lysis..
I would like to know if someone can help me to interpret a fatty acid profile from 2 bacteria. They are two different strains from the same species. So I did a Fames analysis with those 2 bacteria and the results that came out show the same percentage of saturated and unsaturated fatty acids between the 2 bacteria, but when I do the sum of each group of fatty acid (C14, C15, C16, C17, C18) I can see that the percentage of each group can be really different, by 20-35% between the 2 bacteria.
Can someone help me to find a significance/interpretation of the differences?
Your question touches on the unresolved issue of what is the boundary of a species (remember there is no general universally agreed definition), so two isolates can be quite different while classified being blonging to the same species, e.g., by 16S rRNA sequence; I have observed with strains having above 99% similar 16S rRNA sequence similarity quite different amount of lipds as well as cell wall) Different strains can have different total amounts of lipids while the ratio is the same. Small differences in the regulation of lipid anabolism/metabolism can change significantly the total amount of lipids. Taking in account the variability of the lipids by growth conditions -- some times minor differences, such as cell size different doubling times so the growth phase is different etc.)
In short: I would not put too much emphasis on your observation, except the lipids play a major role in your research project. In that case: First I would carefully repeat the experiment, i.e., lipid analysis under strictly controlled growth conditions ( e.g., same protein content / cell numbers at harvesting, check microscopically size and morphology while doing a growth curve, etc.)
Recently my Rhizobium culture has a thick and viscous solid that is some times as much as half the volume of the sample. I have tried to plate the culture but there is no obvious contamination and the EMP produced seems to be stable. When I try to make cell pellets, they will not stick to the wall of any tube. Increasing and decreasing centrifugation does not help.
Can some one help me to figure out what is going on with my Rhizobia? This is a unique isolate and I can not get any more.
If you have eliminated the possibility of contamination, then as mentioned above, try different media or cultural conditions. Sometimes, rhizobia clump/agglutinate when grown in liquid culture. I presume that is what you mean when you say you have a thick and viscous solid. I also presume that you do not see this behavior on plates. Clumping/agglutination can be due to a number of causes and many papers have been published addressing this issue.
I agree with Praveen that sequencing from both ends, and making a contig is the best solution. It will give you the longest sequence, and therefore the best phylogenetic information. On the other hand, in most cases, a single read (800 bps) will be enough to nail down the bacterial species. It is only when you are dealing with less characterized bacterial groups or a novel bacterial sequence, that you might need a longer sequence to be able to get an exact answer, or to get a reliable phylogenetic tree placement.
Does anyone have any experience with L-drying bacteria? I'm evaluating this technology and I don't have direct experience with this interesting approach. What is the best temperature and solvent for enterobacteria?
Nosotros hemos trabajado con freeze-drying, encontrando que la trehalosa en concentración 200 mM es buena protectora de bacterias como Pseudomonas putida, E. coli, entre otras. Nunca he trabajado con L-drying, pero algunas veces por las prisas no congelamos las muestras bacterianas y sometimos a vacío directamente, ahí observamos que de cualquier forma la trehalosa y otros azúcares no reductores protegen a las bacterias de la pérdida de agua. Por si te sirve te envío las siguientes referencias:
Muñoz-Rojas J., P. Bernal, E. Duque, P. Godoy, A. Segura and J. L. Ramos. 2006. Involvement of cyclopropane fatty acids in the response of Pseudomonas putida KT2440 to freeze-drying. Applied and Environmental Microbiology 72(1): 472-477. ISSN: 0099-2240.
Morales-García Y. E., Duque E. Rodríguez-Andrade O., de la Torre J., Martínez-Contreras R. D., Pérez-y-Terrón R y Muñoz-Rojas J. 2010. Bacterias preservadas, una fuente importante de recursos biotecnológicos. BioTecnología 14(2):11-29. ISSN: 0188-4786.
Reviewer of my research article asked about the distribution (either normal or not normal) of some values including the growth associated and non growth associated constants for bacterial growth. These are constants obtained by plotting the growth rate and production rate against each other.
SInce your inserts are not very large ..there should be any problem....Take the molar ratio of vector and insert into consideration when doing the ligations...If you want to be sure about ligation...you can run the ligated sample on an agarose gel along with unligated sample side by side...and visualize it
My bacterial strain was found to be positive for AHL production using CV026 as a biosensor strain and gave big zones of purple coloration, but no peaks for AHL were detected on LC-MS analysis. I can't understand the reason. Extraction was done with ethyl acetate as well as with dichloromethane, at pH 2 and also without any change in pH.
Most probably you are not detecting AHL with CV026. Biosensor strains are highly sensitive but sometimes are also not very specific. Maybe you have an unspecific (false positive) result with CV026. Several articles report that biosensors detect compounds from the culture medium not related to AHLs.
I want to assess the effect a certain compound in the macromolecular biosynthesis of bacteria, for that I intended to use tritiated-thymidine and uridine labelled with phosphorus-32. Can I add both isotopes at the same sample and measure the radiation by liquid scintillation counting?
Yes, of course. The beta energies of these two radioisotopes differ considerably. 3-H emits the lowest, 32-P the highest energy of tracers used in biology. Your instrument will almost certainly be able to measure in different energy windows. The manual will tell you more. In any case you are always able to measure 32-P completely separated from 3H (high energy window), whereas it might be possible to detect some 32P in the 3-H (low energy window), too. This depends on quenching behaviour of your samples. Quenching substances that shift the energy to lower channels and thus affect your counting results are: water and phenol and many coloured substances. The best sample is dried and bound to a lass fiber or nitrocellulose filter. In any case: All samples in one experiment must have the same composition. So, if there is water in your samples, make sure it's the same amount in all vials. It is also helpful to have the same amout of cell extract in all vials.
Please read the manual of your conter for settings, and you will see, with some easy precautions, things are far from being difficult.
I have selected a powerful new compound, who can kill some G positive bacteria at a low concentration. Now, I have only several grams of this compound, and I want to figure out the possible target of this compound by selecting resistant mutants. I have some experience on selecting resistant mutants on a plate and I know that needs more compound, so, who can provide some selection method consuming less compound, such as selecting in broth?
When I precipitate the bacteria for making the competent cells, the bacterial pellets are somewhat black. I use LB medium(10+10+5) & DH5-alpha for precipitation. I centrifuge them at 8000 rpm for 10 minutes. I'm sure that they aren't contaminated because when I use them for transformation, they work properly.
We are beginning to work with a group of students to develop a database for us to track our samples through our very complicated analysis process. We are thinking about using Microsoft SQL server. Do you have any advice on database construction/management/usage? I have no experience with databases, so I appreciate any advice you can give me.
I would suggest that in a first step you have to define all lab processes which are supposed to be tracked in your new database. This will probably be quite a lot of work (depending on the size of the lab) as you might have to talk to a few different people to ensure that you fully understand what everyone is doing in their particular workflow. This information will then form the basis to generate a high level flow chart depicting how a sample will be handled throughout its processing (all the different possible assays, experiments, sending to other providers), as well as what kind of data it will produce and where this data ends up at the moment. The chart will probably be very complex, but it will achieve several things:
1) You will have a much clearer picture of what kind of processes and dataflows are happening in the lab
2) this will form the basis for all further discussions and planning of the software implementation
3) it will also form the basis of what exactly needs to be implemented as a minimum requirement in order to have project milestones
4) it will allow the generation of test cases to later test the functionality of the database
5) the IT guys love flowcharts and it will make it much easier for them to design a database which can handle all the processes
These steps should come well ahead before making a decision which kind of DB system or web app environment should be used to implement the solution. All the underlying logic needs to be developed before anyone starts producing code.
However, before starting to develop some custom database and frontend I would also suggest to look at what is currently available and if existing solutions could be customized to your requirements. E.g. http://www.labkey.com/ is a free solution for managing biomedical data (both research & diagnostic) but it is not trivial to set up and customize - on the plus side it might save a lot of work reinventing things that are already out there.
I would start with a literature search. The obvious (classic) way of doing this is to use bacterial lysates to coat the plates, block, incubate diluted serum in blocking solutions and the detect the bound antibodies with a suitable conjugate. A lot of individual details relate to the conditions that you have in your lab.
The problem of using bacterial lysates, however, is that they contain biomolecules (proteins, LPS, lipids etc.) that have a high similarity to other bacteria and lead to cross-reactivity. So, the clinical specificity of the test is not very high. Typically, today one would use recombinant proteins, or even fragments, because they make possible to detect only specific antibodies. Or you employ extraction protocols to end-up with specific fractions. Often it is a good idea to enrich surface proteins but it is not always sufficient That's why you need to do the literature search in the first place. Hope to find ideas about the antigens to employ.
I would love any suggestions you have for practical ways to avoid contaminating PCR reactions. What are some simple things to do? I know the basics, but I keep having sporadic contamination, so I would love some new tips. What do you do to keep your PCRs clean?
I have obtained a few compounds from bacteria which could only be dissolved in DMSO. Since DMSO has a high boiling point, I am finding it difficult to crystallize or solidify the compounds for further analysis. It would be helpful if there were some simple methodologies to completely evaporate or to separate the compounds from DMSO, e.g. partitioning using solvents.
I don't know what you mean by "only soluble in DMSO" but I think that your compounds are probably able to dissolve in organic solvents. For example, if the compounds are soluble in chloroform, you can remove DMSO by water extraction (3 washes with water for example) then you can remove chloroform by evaporation (a stream of nitrogen for example) and finally sublimate the traces of DMSO with freeze-drying. Then the compound can be again dissolved in chloroform for further use.
I want to take samples of bacteria that have been grown in media and amplify the 16S rRNA gene without doing a DNA extraction. I am using SYBR and amplifying a 200 bp fragment. I have done a bunch of serial dilutions, and once I dilute the sample to a certain point, (about 10x) I start getting amplification. I am looking for advice on optimizing my reaction, and things to watch out for. It is a diverse community, and I am not sure what species are there (the source was a fecal sample).
It can be done, but the end result is not going to be very relevant.
First, growing your sample in a medium will already limit the species that you will find after culturing, and the abundance ratios will probably already be very different from those in stool. Certain bacteria will grow faster than others, depending on the type of medium you choose, so the ratios of those bacteria will be higher than in the original sample.
Second, if you just do a PCR on grown bacteria, without DNA purification (but maybe just a simple boiling step), you will only detect the bacteria that are very easy to lyse. The first PCR step is a near-boiling step, so you will break open some bacterial species, but many others are harder to lyse and won't be detected.
So you could do a qPCR and get results, but the results will be very different from the abundances in the original sample. Even if you do a qPCR on a gene that all bacteria have, such as the 16S rRNA gene, the number of bacteria (or rather 16S copies) is not going to match that of the original sample. I am not sure how that would be relevant, but maybe it makes sense in your specific application.
Anyway, I would highly recommend skipping the culturing step and doing a DNA purification directly on the stool sample. The results from a qPCR on that DNA would be much more relevant to the real abundances in the original sample.
I have seen several papers recently in which plate (viable) counts of bacteria were done on samples which had been frozen in liquid nitrogen and stored at -80 for weeks to months. Does someone know of a reference documenting this procedure?
I work in very big pots of 150L and I want to follow the distribution of bacteria overtime. The objective is to understand bacteria movement into the soil under different conditions (soil type and watering intensity). So I try to develop non-destructive sampling methods I could use overtime to check the presence of bacteria into the soil. I heard about methods like pushpoint sampling but I was wondering if there were others methods or tools which already exist?
I was asked to generate and analyse the phylogeny of NDM-1 protein sequences for my bioinformatics project, but as NDM-1 protein- coding gene is the same in different Enterobacteriaceae it didn't generate a meaningful phylogeny. I was then asked to research and show the phylogeny of different NDMs classes, I am not sure yet whether that would give me a useful phylogeny.
This is normal that you found several identical NDM-1 sequences because in fact many persons have deposited in Genbank the same sequence so-called NDM-1. That's why you have to use only one sequence of each variant (almost 10 variants now).
Concerning the phylogeny, you can but since all the variants are just NDM-1 with one or two ponctual substitutions, the result of phylogeny will be difficult to interpret.
It's so easy to prepare SL7 or SL8. Just follow the original paper. All the salts are Standard and are certainly present in your Institution - maybe with a Little help from the Institute of Inorganic Chemistry. Courage!
When I grow the colonies in R2A medium, I saw clear halos surrounding the colonies. The medium is prepared with seawater and maybe some salts are precipitated after sterilization. Could this be related with the halos?.
It is important for us to know what type of bacteria you cultured in R2A saline medium. It could be that the bacterial strains you are talking about came from seawater and the intestines of benthonic organisms. Cultivable bacteria could be characterized and classified as either psychrotolerant or psychrophiles. Presence of halos around bacterial colonies when they grow in mediums, such as R2A saline medium, could be attributed to protease production. Studies show that more protease-producing strains (around 67%) are detected from the isolated bacteria coming from benthic invertebrates as compared to those obtained from seawater (around 33%), with protease production under neutral conditions resulting in milk protein hydrolysis halos between 27 and 30 ± 2 mm in diameter (2011 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim).
I think the Carba-NP assay is the most convenient one!
See also the following papers:
Vrioni G et al. Comparative Evaluation of a Prototype Chromogenic Medium (ChromID CARBA) for Detecting Carbapenemase-Producing Enterobacteriaceae in Surveillance Rectal Swabs. J. Clin. Microbiol. June 2012 vol. 50 no. 6 1841-184.
Huang TD et al. Comparative evaluation of two chromogenic tests for rapid detection of carbapenemase in Enterobacteriaceae and in Pseudomonas aeruginosa isolates. J Clin Microbiol. 2014 Aug;52(8):3060-3.
Dortet L et al. CarbAcineto NP test for rapid detection of carbapenemase-producing Acinetobacter spp. J Clin Microbiol. 2014 Jul;52(7):2359-64.
Nordmann P et al. Rapid detection of carbapenemase-producing Enterobacteriaceae. Emerg Infect Dis. 2012 Sep;18(9):1503-7.
I was growing my Arcobacter happily using the recipe for Vandamme media (broth and agar). Then suddenly I had either poor growth or not growth in VD agar. All the ingredients were the same except the agar which I purchased a cheaper one. Could this be reason?
What is the best way to quantify the toxicity of short chain alcohols, poly alcohols and organic acids for bacteria? I have been working on production of some of these compounds and the level of toxicity to bacteria is frequently asked by reviewers.
The best way to quantify product (or substrate) inhibition by such compounds is to modify or introduce a dedicated term in the specific growth rate expression. The mathematical form that describes best the inhibition depends on the compound in question, the strain and the cultivation conditions (can be linear, exponential or other) and must be accessed experimentally. There are a number of papers published on this topic. I myself have worked mostly on inhibition by organic acids (e.g. VELIZAROV, S., BESCHKOV, V. (1998) Biotransformation of glucose to free gluconic acid by Gluconobacter oxydans: substrate and product inhibition situations. Process Biochemistry 33, 5, 527-534. which available on my researchgate page), but conceptually these inhibition models can be applied to any inhibiting compounds. Hope this helps.
I have a problem with the PCR amplification of nickel& cobalt resistance genes. The genes are not getting amplified under normal PCR mastermix and temperature conditions. Is there anything that I have to concentrate very specifically in getting the gene amplified?
Check if your DNA is pure or not. If it is not pure, process it and then do PCR. some impurities may interfere your PCR reaction. and also check whether the primer is specific to that gene or it is universal or not. if it is specific for a particular group of organisms get a universal primer and try.
I want to transform E.coli strain MG1655 with my plasmid (pMALc-2x derivative, MBP fusion). Unfortunately, I cannot get any clones. I can transform those cells with different plasmid (so they are competent) and I can transform my plasmid to different strain. It's just this combination which doesn't work. It's not because of my gene - I don't get any clones even if I try with empty plasmid. Does anyone know what could be the reason?
I have been trying to find bacteria or fungi strains that are resistant to cold pasteurization with electric shocks, or any bacteria that is immune to electroporation or any sort of electric discharge, but I have been unsuccessful in my research. Has anyone heard about any sort or strain like that?
A very interesting question, the answer to which I would be curious to know. Whether an "immortal" strain could exist and survive electric treatment...? The main point is that one way or another (through electroporation or ohmic heating of the medium, if you insist with discharging and increase the electric field strength, any cells will be ultimately destroyed. I have written something about this some years ago. If you are interested to read it, the reference is "Electric and magnetic fields in microbial biotechnology: possibilities, limitations and perspectives." In: Electro- and Magnetobiology 18, 2, 185-212 (it is available on my RG page).
I am studying the growth and acid tolerance of various E. coli strains. I am growing cells in minimal media containing glycerol as a carbon source. During this study, I have noticed that E. coli RR1 has not shown any growth. Does anybody have experience with E. coli RR1 with glycerol? If so, do you have any advice?
RR1 carries a number of mutations. It is auxotrophic for proline and some genotypes list an auxotrophy for leucine as well. So you should try adding those two amino acids to your minimal medium. Also most E. coli K12 strains require vitamin B1 (thiamine) which needs to be added to minimal medium.
It is possible to develop an anti-HIV therapeutic based on bacteria.
Mycobacterium bacterial species could be a model for such an approach as these bacteria can enter immune cells. Perhaps this could be an application for a synthetically engineered bacteria.
The bacteria would be engineered to display HIV receptors bacterial surface. This would enable the bacteria to absorb free HIV viral particles. The same displayed HIV receptors could be designed to govern bacterial attachment and entry into virus infected human cells that which display HIV envelope proteins. The bacteria could then be engineered to kill cells infected with HIV.
A major advantage of your suggestion is that this bacteria based anti-viral therapeutic would be self replicating.
I'm growing E. coli with an empty plasmid (no gene) for the standardization of the growth rate of other constructs containing different recombinant genes. I found that when I start the induction with IPTG, there is some toxicity production that more or less stops the growth of E. coli containing the empty plasmid.
I tried to grow E. coli with the empty plasmid without inducing with IPTG, but the growth rate is lower than in the majority of my constructs containing different recombinant genes. Does anyone have any explanation or any suggestions about how I can grow my positive control?
Yes, the empty plasmid when induced with IPTG seems to express toxic genes for E. coli. so is not possible, at least pET plasmid to use it empty. See this paper:
Over-production of Proteins inEscherichia coli: Mutant Hosts that Allow Synthesis of some Membrane Proteins and Globular Proteins at High Levels
Bruno Miroux, John E. Walker
"Surprisingly, none of the cells containing the ‘‘empty’’ plasmids produced colonies in the presence of IPTG, except for pET 17b, which gave very small colonies, demonstrating that the plasmids themselves are intrinsically toxic to E. coli BL21(DE3) host cells."
During a recent literature search I came across the term "Bacterium aliphaticum". The pubIication is from 1944 (Johnson et al., J. Bacteriol. 47:373). "B. aliphaticum" is probably not a valid species designation anymore. Has it been reclassified? Is it a valid bacterial species at all? Does anybody have more info on "B. aliphaticum"?
Apr 9, 2013
Didn’t find much information about B. aliphaticum classification or the valid name but some of the references say that it is probably a member of genus Pseudomonas.
Didn't get the same OD (optical density) results with spectrophotometer
Apr 3, 2013
I made a measurement with using both cuvette and microplate for the measurement of growth rate at OD 595nm, but they have big differences in the results. I got 0.2 with ELISA microplate reader (I put 200 ul into the well), but with the same sample I got 0.7 by measuring using cuvette.
I think cuvette gives reliable results because the turbidity of the medium was much more than 0.2, but reading with ELISA reader is easier because I have so many samples to measure. Which one should I prefer?
I think the difference you're seeing is due to pathlength. Cuvettes, in general, will have a 1 cm pathlength which means that light has to travel through 1 cm of sample. When you measure a sample in a plate the pathlength will vary with the volume of sample that you put in i.e. 200 ul may mean light is only traveling through 0.2 cm hence less sample encountered and therefore less scattering/abs. Some plate readers have a pathlength correction function and can estimate the OD of your sample at 1cm, if you could use that you might get a comparable value.
I would think this is the most likely reason that you're seeing a difference and infact it is not due to an error or you doing anything wrong.
How to make a bacterial aggregation test?
Mar 27, 2013
I want to make a fast and easy aggregation test for Salmonella species. What would you recommend?
easy and fast ... you could make use of the faster sedimentation rate for aggregates compared to single cells, and determine the optical density decrease in the supernatant fluid, or quantify the biomass in the sediment, or both. You can speed up the sedimentation by a "mild" centrifugation of your sample, like 30 sec at 5.000g. Maybe have a look in our paper PLoS ONE 4(5): e5513 for additional info and inspiration on working with aggregates. Best regards, D
I isolated an Algibacter lectus from my plant tissue, however Algibacter lectus is capable to degrade agar, gelatine and starch. So to confirm that he degrades a specific carbon source, which I investigate during my research, I need a media without agar, gelatine or starch. Algibacter lectus can not degrade Casein. I already cultivating them in liquid with my carbon source and it seems to work, but I would like to confirm, that it is also growing on plates with my carbon source (volatile), but as I said I need to exclude agar, gelantine and starch. So I already know, that it can not degrade casein, so a medium plate with casein would be perfect to show that it can grow on plates but what Casein should I use? I just found the pure protein, which is insoluble in water --> so this does not work and I also want not the really expensive pure protein, I just need Casein. Has anybody experience with Casein as binding agent for plates, which casein to use and how much?
Hi, Eileen, You can use gerlite or phytagel, but both require presence of divalent cations. You can growth also on a tissue paper, or on liquid medium. I am not sure about casein, they contain a lot of proteins, which itself may have a strong effect. I am not sure about PAA (poly-acril-amid)- it may interact with cations in the medium. Alginate may be a good suggestions. Good luck!
There are many types of antifreeze compounds in bacteria, its depend which type of antifreeze you want to isolate viz; protein, amino acid, sugars. Please refer our recent publication on cryotolerance mechanism and some novel compound found in psychrotolerant bacteria to survive in extreme cold condition..............Good luck........
Streak plating is the standard method but it is worth looking at your cells using a microscope first - some bacteria stick together or even grow together in "packets" e.g. Micrococcus are usually in tetrads (4s). In these cases it may be necessary to use a method to brak them apart e.g. Passing them through a fine needle, mild sonication or adding tween. The key thing though is to not have too much microbial biomass on your plate otherwise colonies growing near each other can merge.
Once you have prepared your plate incubate it upside down for a day or two depeanding how quickly they grow then use a sterile loop to pick a single colony into suitable growth medium.
I am looking for some DNA to use as an internal standard when we do sequencing. I need to find a species that has never been found in the human gut, but has a medium GC content. If you know where I can order the DNA from, that is even better. I have been looking for something like Nitrosomonas or Rhodopsuedemonas, but I am having trouble finding a good source. Any suggestions?
I exposed my bacteria to different factors which may affect the expression of a gene. However it has affected the growth of one of my bacteria exposed in a certain stimuli. I can't compare CT values from a medium with a "lot" of bacteria and a medium with "less". I'm thinking on diluting until they will have the same absorbance on the spectrophotometer. But I am not sure if it will be fine. If you have any suggestions, feel free to share. Any help would be greatly appreciated.
I am working on Burkholderia, endosymbiont in the subgenus Crispardisia. I would like to know if it is possible to track the circuit of the bacteria in the plant as it is impossible yet at this stage to isolate it. The bacteria already are in the seeds and propagate while the plant is growing, and will settle in the leaf margins, apical shoot, and axilary bud. So I would like to understand how the bacteria propagate, without having to isolate it first.
One suggestion would be to use immunofluorscence -there must be some property of the bacterium that you could use to detect it. An antibody that crossreacts? A Lectin that has binding to the LPS and not the plant? Even without that you should be able to visualise the bacteria in the plant tissue by microscopy. Unfortunately, any method would be destructive for the plant.
If two identical twins were separated after (a cesarean) birth and raised in different environments, would their gut bacteria have co-evolved in similar ways? Or is the development of microbiota more or less exclusively environmental?
If raised in different environments they will have different gut microbiota, however there will be a certain predesposition towards allowance/or inhibition of particular types of bacteria. One good reason for the different microbiomes comes from the fact that the number of genes expressed on the bacterial end is much larger than whatever genetic predisposition the human genome may be expressing. Also, the gut microbiome composition is "reversible" or rather could be shifted towards different composition with change in the diet/environment.
Definitely yes. The primers have to be designed to anneal with the gene sequence you are looking for. You need not digest the plasmid in most cases. Make sure to run a gel with the plasmid and the amplified DNA later to verify if there would be any dimers or unspecific PCR product/s. Most of the primer design software (e.g. Vector NTI) can give you the right Tm, but you may have to do a temperature gradient if you want a very good product especially if you will use the gene for qPCR or transcription studies.
Take out a sample at every time point and extract total RNA. Submit RNA for sequencing (RNA-Seq). Quantify degradative products using any method you use. In principle you don't have to perform 2 experiments simultaneously. Do one where you grow your culture and quantify degradative products at certain points. Then repeat the exact thing but instead extract RNA at those points and submit for sequencing. Do each one in triplicates and if error is small then you can say that certain genes were expressed when certain biproducts formed.
Depending on the organism and enzyme you are interested in you may try contacting culture collections to be advised on bacteria and yeasts they keep. Many of them offer also searchable online catalogs.
There are just a few links to culture collections offering bacteria and yeast cultures:
To answer this question let’s do a thought experiment. You have two bacterial whole genomes which differ in one base pair. Obviously these genomes are different, so there is certainly a sense in which these are different bacteria. The question now becomes, is the difference significant?
If the difference happened to inactivate the production of a lethal bacterial toxin in one of the two bacteria then the difference could be highly significant. It could mean that the one which produced the toxin was a lethal pathogen while the other was entirely benign. On the other hand, if the difference changed no codon in a housekeeping gene it would mean that there would be no detectable difference in behaviour between these two bacteria.
A restriction digest could pick up the difference, if an enzyme was used whose restriction site was changed by the single difference. But this would certainly not be likely to be the case for any one enzyme, and there could well be no known enzymes which would detect the difference. Sequencing, however, would in principle reveal the difference between the two bacteria.
So the answer to the question is, restriction digestion can be used to identify bacteria only in those cases where you already know a lot about the different bacteria which you want to identify. For anything else you need to sequence, and even that may not tell you what you want to know.
I transformed DH5alpha bacteria 2 months ago. I got some colonies on kanamycin plate. I picked a colony and used plasmid after miniprep. Plasmid worked fine but I forgot to make glycerol stock. Now I am trying to grow bacteria again from same plate which was kept at 4C all the time. They are not growing at all. What could be the reason?
Ehm.. 4ºC.. are you sure about that? Compared to environmental strains, E. coli is unfortunately a bit fragile concerning durability. Cold rooms generally labeled as being at 4ºC are sometimes in fact at 5-6ºC, so the metabolism is not quite dormant. If you're talking of a regular freezer, then it's worse: around 10ºC. Therefore: you should never store an E. coli which you intend to work with again for more than 1 month. 2 months, in my experience is stretching their survival next to the limit. You should have streaked part of the material of that original colony to a new plate with the desired antibiotic (i usually use for example a sterile toothpick or a micropipette tip), and then inoculate the tube for the mini-prep. That way you always insure a backup with enough material (i usually make a grid on my backup plate, so it can fit around 50 clones).
Now.. solutions to your problem. I hope you still have the tube with the plasmid you obtained after that mini-prep. Transform a competent E. coli ASAP with that plasmid. And then, don't forget to make the stock ;).
Good responses above. One additional point: you want to insure that your ammonium salts are not above 0.1 % w/v since a range of bacteria are inhibited at higher conc., some even at 0,1% NH4Cl (roughly, pending on the salt 10-20 mM NH4 ions) level. If you use urea, we usually stay below 0.6% w/v. If you are looking for diversity in environmental samples you need to go to very low substrate concentrations but still keep the C/N ratio around 25 -30. If you use nitrate, you need to be careful with the concentration and the CN ratio, since some bacteria (I know some anaerobes) who will at lower Carbon conc only form nitrate which then kill -at least inhibit-- some of your microorganisms. AT high carbon conc they will go all the way to ammonium (see above comment re pH effect)
Can you describe the selection criteria you would use to isolate a bacterium that was able to degrade benzene under aerobic conditions?
The molecular methods are the rapid way to primary identification of unknown bacterial strain. Firstly you should isolate the 16S RNA then make nucleotide sequencing. After that make matching in NCBA database, it will give you the bacteria which have the same sequence. You can confirm with the biochemical assay
I must say I am somewhat surprised about this general question. Are you a high schook student? Then I am excited that you are interested in this subject..If you ae a University student you could have gotten some more info before (sorry for being a little harsh here)
There are hundreds of publications describing the conditions of growing and isolating and characterizing these extremophiles., original publications and many books chapters. For instance you could Google for the book Handbook of extremophiles. (Ed.-In-Chief K. Horikoshi) Springer Verlag Tokyo and other you will find listed on the web and you find the best how-to-do chapters written by the worlds authorities. As another start Google the question, read up and then please come back with very specific questions. Decide what type of hyperpiezophilic (the correct new term) and hyperthermophiic bacteria you want to grow or isolate --in case you are not a microbiologist-- e.g., aerobic or anaerobic, bacteria or archaea, alkaliphile, acidophile, or neutrophiles, chemolithotrophs, organo- or lithoheterotrophs (what substrates you are thinking about, etc, etc, I guess you will have to google these terms also. Another option is to go to the International Journal of Systematic and Evolution Microbiology (For the newer publications perhaps you have to go to a University library but older papers are available electronically) you find hundreds of examples of description how those bacteria were isolated and characterized.
I'm planning on using fluorescent antibodies on soil microbial communities, then sorting them using FACS. Can someone please give me the benefit of their experience and suggest possible methods that I can use for permeabilizing the cells to the antibody without decreasing cell viability. I wish to culture the microbes following sorting. Thanks
I'm not sure if that kit would allow for subsequent culture of cells post-treatment though. It may not kill/destroy the cells immediately but they will not continue to live after the treatment. As the kit says you must analyse the cells within 18 hours.
Fixing and permeablising cells, from my experience, is permanent for mammalian cells. I've no idea if this would be different for microbes due to their cell wall and plasma membrane structure.
The action of the fixative "freezes" the cell's physical state and the permeablisation treatment punches holes in the membrane to allow antibodies in. I don't think most cells would be able to survive this!
If you're looking to continue growing the cells after analysing them then don't fix them. Mild detergents are often used to permeabise mammalian cells to probe their internal structure. You could try using very low concentrations of such detergents to see if you could gently permeabilise the membrane. I'm not sure if work has been done on how damaged the membrane can be for microbes to survive but it could be a trial and error thing for you there. Triton X-100 is a common lab detergent used for this. Sigma sell it. By low concentration I'd imagine something like 0.1% v/v or below.
Depending on what you're looking for, do you have to use antibodies? There are plenty of dyes out there that are lipid soluble and so pass across membranes freely. That would mean you wouldn't have to permeabilise the cells though it may not be clear what the dye would do to the metabolics of the cells once in there and staining it's target...
We are currently handling a good collection of lactic acid bacteria and yeast isolates associated with various stages of an indigenous fermentation process for production of fermented bamboo shoot. Most of them are fairly identified by ARDRA and rRNA gene sequencing (similarity range 97-99%). However, being an untapped ecological niche which is not explored deeply, we are expecting this niche might harbour new novel species.
I would normally subscribe to some of the suggestions given above and do feel Michael Granitsiotis have provided a great reference. However, I must point out that eventhough 16S will serve as a preliminary taxonomic tool it may not be enough. You see SSU's are great for providing taxonomic context and will even narrow your species to a solid genus. However, 16S and other highly conserve genes can only do so much. You still have the rest of the genome to distinguish you species. Some classical test like metabolic pathways are usefull to distinguish your candidate. In my opinion it would be usefull combine 16s rDNA data with metabolic information. A good place to start is using BioLog. Is a fast and easy way to get some metabolic information. Also, sequence some house keeping genes (recN, recA etc...) as supplemental information. The best option would be to sequence the genome but that requires a magnanimous effort. I hope you able to define the species. Best of luck to you.
Why doesn't protein expression work with an already established plasmid?
Nov 28, 2012
Our lab extensively works with protein overexpression in bacterial systems. Recently, almost 4-5 of my colleagues and I have been facing problems in expression of protein after induction. What previously gave a significant yield of protein, has suddenly stopped giving any yield. The yield is so poor that it is only visible as faint bands in western blots. We've tried changing the stocks of IPTG, antibiotics. People working with different strains of bacteria face the same issue. The only trivial difference is that the filter to milliQ plant providing water for LB media was changed! That hardly qualifies as an issue.
Has anyone else had experiences as such? Does anyone have any suggestions?
I would also suspect phage infection. It dosn't necessary lyse the cells in shaken flask cultures at the initial phase of plague, you will experience it later, when everything is infected heavily (in fermenters you could see lysis already). To decide, you can make a phage test with the supernatant of a culture (without antibiotics) of yours on phage sensitive E. coli cell layer. I just can hope for you that I am not right.
I have about 30 samples of plant tissue that I want to test for the presence of phytoplasma bacteria. I hit some stumbling blocks in terms of classical PCR and CTAB protocols. I have a new protocol for CTAB and am wondering if I would be better served to go ahead and do real-time first and then do classical just to confirm my results. What do you think?
Hi Julia! You need a lot of expertise to establish a good working q-PCR (Real Time). Do you have the expertise? You want to analyse only 30 samples?! Do you know the costs for q-PCR? I would give you the advice against q-PCR without any knowledge and only for analysing 30 samples especially if you have a classical PCR who is working!
I am isolating halophiles from salt pans and studying their capability of metal tolerance. I have used 1M and 2M Nacl in the medium during isolation. I have got at 9 - 10 different types of colonies. Do I need to study all of them and still more or only the most salt tolerant ones amongst these will be enough?
Well, 1-2 M NaCl concentration shows that they are moderate halophiles (which is a good concentration of NaCl to study them). 9-10 is not big number you must study them all, even check out for more if you still have source in 0.5-1 M NaCl concentration. You can make your choice of study by looking into obligate halophiles and facultative halophiles if the number of isolates increases much.
I tried inoculating the V. harveyi in saline and sea water and there was no infection. In all the articles I have read I couldn't find a detailed procedure. I want to establish an effective infection system for shrimps with v.harveyi.
My question relates to the excessive foam formation when auto inductive growth media is shaken at high rpms. Instead of having to buy a new agent, I wonder if there's a common agent that I could use to suppress foaming e.g. by adding silica oil? Any ideas?
Hi Toni, we sometimes use PEG400 (as crystallographers we always have PEGs in stock :o) - try some (1-5) droplets from a standard syringe/canule combination and shake your culture (180-220 rpm), to see, if it works for you. Good luck, Magdalena
I am trying to determine how long it takes for oil to be broken down by bacteria. My research deals with the oil effects on the foraminifera and the studies I am looking for might help address some of my questions regarding bacterial consumption from that of benthic foraminifers.
Currently I am trying to study the localization of a novel bacteria which can only grow inside an amoeba host cell. So far I do not know the genome or plasmid sequence of the bacterium. What I am planning to do is to tag the bacterium with a tracking dye so that I can follow its life cycle through its host, in this case Acanthamoeba polyphaga.
1. Phase-contrast is always a first good step provided bacterial cell size allows it.
2. Heads up: GFP-expressing bacteria obviously vary in GFP expression duration and strength. Thus, what worked well with Salmonella and E. coli may not work too well with Campy related organisms.
3. We tried using live/dead Baclight kits but we had a lot of challenges interpreting the results when it came to cellular viability.
4. For us, FISH worked best but since you don't have the sequence yet this may present some challenges.
5. While I have not used it myself I understand EM works excellently as well.
6. I would assume you're interested in investigating the internalized bacterium's viability as well throughout the life cycle, in which case culture-based techniques after disrupting the A. polyphaga trophozoite membranes were the most useful.
I have some environmental water samples in which DNAs have been extracted for PCR and sequenced for a functional gene. Upon blasting sequences with genbank, there were no good matches. The highest match was bacteria (90+% coverage and 70-80% max identity). How can I differentiate between the production by bacteria and cyanobacteria?
Yes, it corresponds to one gene from multiple species (bacteria and cyanobact). Unfortunately, I do not have bacteria cultures to sequence them for phylogenetic tree but that is a good idea. Am still looking for a simpler method if you ahve any? Just recently, my GC-MS results showed positive for that chemical compound so I am going to work on that for PCR.
Hi Kazuo - My samples are from the environment so there will definitely be photosynthetic organisms in it. Hence I cannot say for sure that the chemicals are produced by cyanobacteria or bacteria by performing PCR for that. Say if PCR shows positive for the photosynthetic gene, all I can say is there are photosynthetic organisms in my sample. It still cannot tell me if that functional gene (that I am looking for) is by bact or cyanobact.
Biological Nitrogen fixing Bacteria can increase or decrease emission of Greenhouse gases?
Dec 3, 2012
What is the influence of BNF Bacteria on emission of GHGs?
The main GHG related to the Nitrogen cycle is nitrous oxide (N2O). Nitrous oxide is naturally present in the atmosphere as part of the Earth's nitrogen cycle, and has a variety of natural sources. However, human activities such as agriculture, fossil fuel combustion, wastewater management, and industrial processes are increasing the amount of N2O in the atmosphere. The impact of 1 pound of N2O on warming the atmosphere is over 300 times that of 1 pound of carbon dioxide (http://epa.gov/climatechange/ghgemissions/gases/n2o.html). Direct emissions of N2O result primarily from microbially driven nitrification and denitrification processes, together with non-biological chemodenitrification. Direct sources include those where N2O is emitted directly to the atmosphere from cultivated soils and fertilized and/or grazed grassland systems. Indirect emissions result from transport of N from agricultural systems into ground and surface waters through drainage and surface runoff, or emission as ammonia or nitrogen oxides and deposition elsewhere, causing N2O production. In this regard, biological nitrogen fixation and in particular industrial nitrogen fixation increase N inputs available for processes such as nitrification and denitrification, with a potential positive feedback on the production of N2O.
pH of crude bacteriocin is near 4.5. But optimum pH for activity of Proteinase K is 8. For treatment pH of crude bacteriocin was set to 8 and after treatment reversed back to 4. But activity remains the same. The same happens with pepsin, trysin.
To check if you have a bacteriocin you can try heat-inactivation (10 minutes boiling or even 30 minutes of autoclaving for heat-resistant ones). Do a control for proteolytic activity of your enzymes that you can check on SDS-PAGE (BSA, or even a simple molecular weight marker can serve as a control for digestion) add a test protein to an aliquot of your sample, perform a digestion and run a gel.
It is believed that mitochondria have evolved from bacterial systems. We all knew that mitochondria have a very precise electron transport chain system and have a clear cut pathway to maintain chemi-osmotic gradients and to yield energy molecules. If all organisms could produce energy precisely, then it would be easier to sustain life. What I understand is that the objective of evolution is to adapt something in order to sustain life on the earth. Then why did bacteria not evolve with their own mitochondrial systems?
The conversation is a bit confusing but maybe this will help a bit (1) optimizing *efficiency* of energy production is not the single most important factor driving evolution of single cell organisms or functionality of cells in complex organisms. Efficiency requires complexity and additional energy expenditure. Sometimes all you need is a certain level of efficiency. If all you do is multiply in a reasonably nutrient rich environment, why bother with unneeded genetic complexity? (2) mitochondria vary from organism to organism and tissue to tissue. They evolved to adapt to their cell environment over hundreds of millions of years. Some are more efficient than others. ATP production may not always be their prime or most demanding function under certain circumstances. Stem cells are glycolytic. Adipose tissue during hibernation are uncoupled. Etc. Aerobic bacteria do just fine in their environments. Protoeukaryotic host of protomitochondria was probably more concerned with detoxifying O2 than having more ATP. That was a bonus
I am working on a GST pulldown from BL21 cells. I lyse the cells in PBS, 1% triton, DTT, and protease inhibitors with sonication. After I pull down one of my fusion proteins over night at 4 degrees, the beads incubated with my fusion protein which is an additional 300 amino acids including a RING domain and possibly unstructured region, clump together very strongly. I have to pipet up and down very hard to break the clump which is obviously not ideal. The RING domain binds Zinc - I grow the bacteria with additional metals and have emoved the EDTA from my washes but this hasn't solved the clumping problem. Any ideas?
Your protein is clearly aggregating. The aggregation is so strong that the beads are clumped. Take a look at the unstructured region... sometimes these regions are strong interactors. You can try to modify slightly the pH for the binding or increase the ionic strenght to decrease polar interactions.
In a separate experiment you can try also to add glycerol at 5-10% to see if the aggregation is caused by hydrophobic interactions.
in general terms speciation of Pseudomonas can be done based on pigment it produces. Kings media can be used
Bacterial adhesion analysed by quartz crystal microbalance (QCM, QCM-D)
Dec 5, 2012
We observed both frequency decrease and increase during bacterial adhesion in QCM flow systems, dependent on the substrate surface state, the bacteria, and the applied QCM-system. Does anyone have any possible explanations?
I am developing a method to measure the number of cells in a sample from an anaerobic culture using qPCR, and I want to use gDNA as a standard. I am wondering if anybody has any advice about how to do this. I have been doing serial dilutions of both cells and gDNA, and I am getting okay results. I think I can calculate the amount of DNA per cell, figure out how many cells are in each well, figure out the 16S copy number, and compare it to the DNA. I am using e coli right now. I appreciate any advice about this.
Have you found that your gDNA curve and your cell-derived DNA curves have different slopes? If so, there is a mathematical way to reconcile the 2 universes and calculate copy number of the cell gDNA. Hopefully the operon number is the same for 16S in both the gDNA standard and the e coli? Or, are you also analyzing an assumed "single-copy" gene here somewhere?
If you need to 'reconcile the 2 universes' mentioned above - the attached has worked in these situations.