SEA-PHAGES | All posts created by viknesh

Link to this post \| posted 14 Apr, 2021 19:25
viknesh	kmaclea I wanted to send an update that we have conclusively proven the source of our contamination giving us so many problems. The agar we were using to make the top agar appears to have been contaminated with something that could survive 40 minutes of autoclaving (volume 500 ml). Now that we have switched out the agar, our contamination issues have gone away completely. Just an update for anyone following the thread. Kyle Thrilled to hear it, Kyle. Hope your students had fun figuring this out.

Link to this post | posted 14 Apr, 2021 19:25

kmaclea
I wanted to send an update that we have conclusively proven the source of our contamination giving us so many problems.

The agar we were using to make the top agar appears to have been contaminated with something that could survive 40 minutes of autoclaving (volume 500 ml).

Now that we have switched out the agar, our contamination issues have gone away completely.

Just an update for anyone following the thread.

Kyle

Thrilled to hear it, Kyle. Hope your students had fun figuring this out.

Posted in: Phage Discovery/Isolation → Contamination of top agar and possibly other components

Link to this post \| posted 03 Apr, 2021 16:57
viknesh	kmaclea So here's an update. https://docs.google.com/document/d/1294ZwYilXMbQA6n0eFrWouodmvBk1M4ZOL4AsPiUsX8/ Thanks for sharing all this, Kyle. I can't say for sure, of course, but it looks like you might have multiple things going on. First - the media. Did you try imaging the most recent cloudy bottle of top agar (the one that your TA added calcium and dextrose before authoclaving?). I ask because CaCl2 precipitates when heated for prolonged preiods (i.e. autoclaving), and the amount of precipitation depends on the pH and other salts in solution. While I am not discounting your contamination (all those beautiful microscopy images), the cloudiness in the recent bottles might just be precipitate, and cloudy top agar may be sufficient for plaques to look cloudy - in which case your contamination problems might be isolated events and not a widespread problem. Second - I think those contaminated spots on the enriched plates are not coming from the top agar, but instead from the samples spotted themselves. This is fairly common when the enrichment is not properly filtered, typically because there has been a rupture in the filter membrane. I used to see this fairly often and so now buy the fairly "tough" GDX Whatmann filters that rupture less often (but which still occasionally do). Of course, maybe you have mycoplasma, which can pas through the filter). For your plaque assay plates that are very clearly contaminated, again, this might be a problem with the sample itself being contaminated since some other plates look great - are these also filtered soil samples? If you performed/perform a dilution series with these samples, my guess if that you'll see the contamination dilute out too. Kyle, regardless of what's happening, the fact hat you're engaging your students in trying to figure out this issue is pretty awesome! What a ride for them Vic Edited 03 Apr, 2021 16:59

Link to this post | posted 03 Apr, 2021 16:57

viknesh

kmaclea
So here's an update.
https://docs.google.com/document/d/1294ZwYilXMbQA6n0eFrWouodmvBk1M4ZOL4AsPiUsX8/

Thanks for sharing all this, Kyle. I can't say for sure, of course, but it looks like you might have multiple things going on.

First - the media. Did you try imaging the most recent cloudy bottle of top agar (the one that your TA added calcium and dextrose before authoclaving?). I ask because CaCl2 precipitates when heated for prolonged preiods (i.e. autoclaving), and the amount of precipitation depends on the pH and other salts in solution. While I am not discounting your contamination (all those beautiful microscopy images), the cloudiness in the recent bottles might just be precipitate, and cloudy top agar may be sufficient for plaques to look cloudy - in which case your contamination problems might be isolated events and not a widespread problem.

Second - I think those contaminated spots on the enriched plates are not coming from the top agar, but instead from the samples spotted themselves. This is fairly common when the enrichment is not properly filtered, typically because there has been a rupture in the filter membrane. I used to see this fairly often and so now buy the fairly "tough" GDX Whatmann filters that rupture less often (but which still occasionally do). Of course, maybe you have mycoplasma, which can pas through the filter).

For your plaque assay plates that are very clearly contaminated, again, this might be a problem with the sample itself being contaminated since some other plates look great - are these also filtered soil samples? If you performed/perform a dilution series with these samples, my guess if that you'll see the contamination dilute out too.

Kyle, regardless of what's happening, the fact hat you're engaging your students in trying to figure out this issue is pretty awesome! What a ride for them smile

Vic

Edited 03 Apr, 2021 16:59

Posted in: Phage Discovery/Isolation → Contamination of top agar and possibly other components

Link to this post \| posted 31 Mar, 2021 11:52
viknesh	kmaclea So, this is going to sound insane, but we've been running into recurrent bacterial contamination that a couple of attempts to fix doesn't seem to be fixing. We are trying to use both A. globiformis and M. foliorum for phage discovery. We had a lot of luck getting plaques originally, then at some point most of our students' phages have been unable to propagate further (no plaques). We've done countless direct and enriched isolations. A few weeks ago we had a contamination issue and re-made components. Things got better for a time, but then we were back in problems again. Last week there was obvious rampant contamination again. We received new glycerol stocks for bacteria, remade all the PYCa, all the top agar, all the dextrose, CaCl2, and CHX. The last three were sterile filtered, the first two were autoclaved. Autoclaving was standardly done with 500 ml at a time and 45 minute autoclaving at 121C. By all indications, other media being made using the same autoclave is just fine although no biological testing of the autoclave has been done recently. Most recent tests seem to indicate it is the top agar in which we are seeing contamination. Even in freshly made top agar, freshly autoclaved, with the other components all sterile filtered in the hood. Attention being paid to sterile technique during the handling, in the hood. What could be making our top agar contaminated? We have been using a metal bead bath for incubation of the top agar, but we are seeing this even in freshly made top agar. Beads in the bead bath have also been recently decontaminated. Ideas of any kind would be most definitely welcomed! Kyle Hi Kyle, Looks like there ate at least two problems: 1) not being abe to propoagate a phage and 2) contaminated materials. For #1, are you seeing this for both hosts, or just one? If the latter, which one. Also, will the control phage also not plaque? For #2, are you able to share images of the contamination? I can;t tell if the top agar is getting contaminated as you use it or whether it was never actually fully sterilized. For example, what happens to the freshly autoclaved top agar, still molten in the bottle with the lid closed, if left in the beadbath for a few days - never opened? Do you see microbial growth in the bottle, or only when you use it? Vic

Link to this post | posted 31 Mar, 2021 11:52

viknesh

kmaclea
So, this is going to sound insane, but we've been running into recurrent bacterial contamination that a couple of attempts to fix doesn't seem to be fixing.

We are trying to use both A. globiformis and M. foliorum for phage discovery. We had a lot of luck getting plaques originally, then at some point most of our students' phages have been unable to propagate further (no plaques). We've done countless direct and enriched isolations.

A few weeks ago we had a contamination issue and re-made components. Things got better for a time, but then we were back in problems again. Last week there was obvious rampant contamination again. We received new glycerol stocks for bacteria, remade all the PYCa, all the top agar, all the dextrose, CaCl2, and CHX. The last three were sterile filtered, the first two were autoclaved. Autoclaving was standardly done with 500 ml at a time and 45 minute autoclaving at 121C. By all indications, other media being made using the same autoclave is just fine although no biological testing of the autoclave has been done recently.

Most recent tests seem to indicate it is the top agar in which we are seeing contamination. Even in freshly made top agar, freshly autoclaved, with the other components all sterile filtered in the hood. Attention being paid to sterile technique during the handling, in the hood.

What could be making our top agar contaminated? We have been using a metal bead bath for incubation of the top agar, but we are seeing this even in freshly made top agar. Beads in the bead bath have also been recently decontaminated.

Ideas of any kind would be most definitely welcomed!

Kyle

Hi Kyle,

Looks like there ate at least two problems:
1) not being abe to propoagate a phage and
2) contaminated materials.

For #1, are you seeing this for both hosts, or just one? If the latter, which one. Also, will the control phage also not plaque?
For #2, are you able to share images of the contamination? I can;t tell if the top agar is getting contaminated as you use it or whether it was never actually fully sterilized. For example, what happens to the freshly autoclaved top agar, still molten in the bottle with the lid closed, if left in the beadbath for a few days - never opened? Do you see microbial growth in the bottle, or only when you use it?

Vic

Posted in: Phage Discovery/Isolation → Contamination of top agar and possibly other components

Link to this post \| posted 08 Feb, 2021 01:21
viknesh	kmaclea Dear Colleagues We are running phage discovery in the winter for the first time. Our first go-round we have a lot of success with direct isolations out of the box. This might have been an outlier leading us astray, but over the last couple of weeks we only had one successful run with direct isolation. We have started enriched isolation but still wondering if we can do better with our direct isolation or if winter in New Hampshire is just going to be more challenging. Anyone have thoughts? Kyle Hi Kyle, From what I can tell based on findings from across the SEA, the success rate for direct isolation is generally low and inconsistent (unless you keep going back to the same/similar spot). I'll let faculty from the northern reaches of the country share their advice, but I know several who gather soil early (before the ground freezes, and store them in sealed ziplocks at 4C) or use soil from indoor pots. I personally have had success with both methods when isolating phage for M. foliorum, but only by enriched isolation. Good luck!

Link to this post | posted 08 Feb, 2021 01:21

viknesh

kmaclea
Dear Colleagues

We are running phage discovery in the winter for the first time. Our first go-round we have a lot of success with direct isolations out of the box. This might have been an outlier leading us astray, but over the last couple of weeks we only had one successful run with direct isolation. We have started enriched isolation but still wondering if we can do better with our direct isolation or if winter in New Hampshire is just going to be more challenging.

Anyone have thoughts?

Kyle

Hi Kyle,

From what I can tell based on findings from across the SEA, the success rate for direct isolation is generally low and inconsistent (unless you keep going back to the same/similar spot). I'll let faculty from the northern reaches of the country share their advice, but I know several who gather soil early (before the ground freezes, and store them in sealed ziplocks at 4C) or use soil from indoor pots. I personally have had success with both methods when isolating phage for M. foliorum, but only by enriched isolation.

Good luck!

Posted in: Phage Discovery/Isolation → Phage isolation in winter--any differences?

Link to this post \| posted 05 Feb, 2021 01:34
viknesh	cheryl.brown@mnstate.edu I have another PYCa media question. Does the type of peptone you get affect the media? We have been using VWR 97064-330, but there is a non-animal, defatted soy peptone (97064-186) for less than half the price. Has anyone used this? Does the source of the proteins matter in this application? Hi Cheryl, This is a little tricky to answer. While peptones are generally composed of the same components (amino acids, peptides, carbs, salts, vitamins), the ratio of the components and pH of the peptone will vary based on the source material and the process used to generate the peptone. These diffrences have been shown to dramatically impact growth and the behaviour of some cell lines and bacteria but not others, so most people tend to stick with what they know to work. I'm hoping the Hatfull lab will chime in here, but I for one use the recommended peptone because I've not tested any other sources of peptones. If you are keen to give the soy-based peptone a try, then I would advise purchasing a small batch to do some growth comparisons. If you do, I would love to know what you discover. Vic Edited 05 Feb, 2021 01:34

Link to this post | posted 05 Feb, 2021 01:34

viknesh

cheryl.brown@mnstate.edu
I have another PYCa media question. Does the type of peptone you get affect the media? We have been using VWR 97064-330, but there is a non-animal, defatted soy peptone (97064-186) for less than half the price. Has anyone used this?

Does the source of the proteins matter in this application?

Hi Cheryl,

This is a little tricky to answer. While peptones are generally composed of the same components (amino acids, peptides, carbs, salts, vitamins), the ratio of the components and pH of the peptone will vary based on the source material and the process used to generate the peptone. These diffrences have been shown to dramatically impact growth and the behaviour of some cell lines and bacteria but not others, so most people tend to stick with what they know to work. I'm hoping the Hatfull lab will chime in here, but I for one use the recommended peptone because I've not tested any other sources of peptones.

If you are keen to give the soy-based peptone a try, then I would advise purchasing a small batch to do some growth comparisons. If you do, I would love to know what you discover.

Vic

Edited 05 Feb, 2021 01:34

Posted in: Phage Discovery/Isolation → Peptone question

Link to this post \| posted 22 Jan, 2021 21:03
viknesh	cheryl.brown@mnstate.edu We have had some issues with our PYCa agar getting dark while being held at 45-50 C. I am trying to figure out if it could be one of the ingredients, as we did buy new yeast extract and peptone, or if this is normal and harmless. Hi Cheryl, At high temperatures, dextrose and amino acids can undergo the Maillard reaction, which results in a darkening of the media. The products of the Maillard reaction impacts the growth of some bacteria more than others. It may be fine for you to use the darkened media, but it may impact reproducibility of bacterial growth on your plates. The way to reduce the Maillard reaction is to keep dextrose and amino acids in as cool a solution as possible. This means waiting for freshly autoclaved/molten PYCa agar to cool to ~ 60C before adding dextrose, minimizing the number of time you reheat (e.g. in the microwave) PYCa agar once dextrose has been added, and not keeping molten dextrose in the warm bath for extended periods of time. In our lab, we add dextrose to molten PYCa agar at ~ 60C, then allow the media to solidify for storage. When we want to pour plates, we melt the solidified agar in the microwave, allow it to cool to 60C before adding supplements like cycloheximide, then pour the plates. We don't keep PYCa agar in the molten state for long. I hope this is helpful.

Link to this post | posted 22 Jan, 2021 21:03

viknesh

cheryl.brown@mnstate.edu
We have had some issues with our PYCa agar getting dark while being held at 45-50 C. I am trying to figure out if it could be one of the ingredients, as we did buy new yeast extract and peptone, or if this is normal and harmless.

Hi Cheryl,

At high temperatures, dextrose and amino acids can undergo the Maillard reaction, which results in a darkening of the media. The products of the Maillard reaction impacts the growth of some bacteria more than others. It may be fine for you to use the darkened media, but it may impact reproducibility of bacterial growth on your plates.

The way to reduce the Maillard reaction is to keep dextrose and amino acids in as cool a solution as possible. This means waiting for freshly autoclaved/molten PYCa agar to cool to ~ 60C before adding dextrose, minimizing the number of time you reheat (e.g. in the microwave) PYCa agar once dextrose has been added, and not keeping molten dextrose in the warm bath for extended periods of time.

In our lab, we add dextrose to molten PYCa agar at ~ 60C, then allow the media to solidify for storage. When we want to pour plates, we melt the solidified agar in the microwave, allow it to cool to 60C before adding supplements like cycloheximide, then pour the plates. We don't keep PYCa agar in the molten state for long.

I hope this is helpful.

Posted in: Phage Discovery/Isolation → PYCa media turning dark

Link to this post \| posted 10 Dec, 2020 18:23
viknesh	Hi Maria We have imaged phage using the TEM mode on an SEM. The quality and magnification are understandably lower. I've attached one of the better micrographs, and you can see that a lot of the usual detail we can visualize with TEM is lost. 1.64Mb

Posted in: Phage Discovery/Isolation → Electron Microscopes

Link to this post \| posted 19 Nov, 2020 18:14
viknesh	erin.windsor The only enzyme that cuts well for us in Gordonia rubripertincta is NspI. See attached. Thanks for sharing this, Erin. We too have found that NspI reliably cuts DNA of phages isolated on G. rubripertincta. We've also found that BamHI, HindIII, and SacII also cut a few of the G. rubripertincta phages we've isolated. Your results will defintiely help us get a better grasp of which enzymes are best to recommend moving forward. Vic

Link to this post | posted 19 Nov, 2020 18:14

viknesh

erin.windsor
The only enzyme that cuts well for us in Gordonia rubripertincta is NspI. See attached.

Thanks for sharing this, Erin. We too have found that NspI reliably cuts DNA of phages isolated on G. rubripertincta. We've also found that BamHI, HindIII, and SacII also cut a few of the G. rubripertincta phages we've isolated. Your results will defintiely help us get a better grasp of which enzymes are best to recommend moving forward.

Vic

Posted in: Gordonia → Gordonia DNA not cutting

Link to this post \| posted 19 Nov, 2020 16:58
viknesh	Hi Amy, Would you mind sharing a little bit more information about what you're using and seeing? For example: What restriction enzymes are you using? Did you also include DNA for your control phage? If so, what phage is it? Chidiebere? Could you share a gel image of what you are seeing? Thanks. Vic asprenkle Anyone have any trouble getting a good RE digest of Gordonia DNA? I've got good DNA from M. foliorum, and we're trying Gordonia rubripertincta for the first time this year. I've noticed it's a bit waxier than M. foliorum. I'm getting OK DNA concentrations with a new Wizard kit, but my Mf DNA cuts where my Gr does not. Any tips/tricks I'm missing? Thanks!

Link to this post | posted 19 Nov, 2020 16:58

viknesh

Hi Amy,

Would you mind sharing a little bit more information about what you're using and seeing? For example:

What restriction enzymes are you using? Did you also include DNA for your control phage? If so, what phage is it? Chidiebere? Could you share a gel image of what you are seeing?
Thanks.
Vic

asprenkle
Anyone have any trouble getting a good RE digest of Gordonia DNA? I've got good DNA from M. foliorum, and we're trying Gordonia rubripertincta for the first time this year. I've noticed it's a bit waxier than M. foliorum. I'm getting OK DNA concentrations with a new Wizard kit, but my Mf DNA cuts where my Gr does not. Any tips/tricks I'm missing?
Thanks!

Posted in: Gordonia → Gordonia DNA not cutting

Link to this post \| posted 21 Feb, 2020 20:51
viknesh	Hi Sarah, It's likely that the liquid culture was contaminated. As such, your lawn is composed of more than one bacterium - M. foliorum and other bacteria. Phages will still be able to infect M. foliorum, but plaques may not be apparent because contaminating bacteria, which cannot be infected by those same phages) are able to continue to grow evenly on the plate and therefore also within the plaque. The weird smell is likely due to the contaminant. Hope this helps. Vic Thiel Hello, I think that we are having a similar problem to the post above that says "D29 plaques barely visible on M. smeg" but I'm not sure. We are doing the phage isolation for the 2nd semester (so I'm not very experienced) and the students were on their 2nd round of purifications and starting to do the 3rd round and then make webbed plates. Our host is M. foliorum. I take new M. foliorum out of the -80C every 2 weeks, streak a plate and then shake it to make new stocks for them to use. The entire class started getting very similar looking plaques, that were difficult to see and smelled weird. (See image). The plates labeled "old" bacteria is what we had been using. The plates labeled "new" bacteria was when we picked a new colony from the same streak plate and it starting giving us the odd results. The difference in the incubation times was because we thought if we gave the 10^-3 plate a little more time- it would fully lyse, but it didn't. We picked a another new colony from the M. foliorum streak plate, grew it up and it seems to have fixed the problem (no pic). Just in case, I made a new M. foliorum streak plate from a different glycerol stock and will be growing that up for future use for the students. So we might have fixed the problem, but what do you think happened? Did we get lucky and pick a mutant colony? Any insight would be awesome. I'll let you know if the problem is not fixed.

Link to this post | posted 21 Feb, 2020 20:51

viknesh

Hi Sarah,

It's likely that the liquid culture was contaminated. As such, your lawn is composed of more than one bacterium - M. foliorum and other bacteria. Phages will still be able to infect M. foliorum, but plaques may not be apparent because contaminating bacteria, which cannot be infected by those same phages) are able to continue to grow evenly on the plate and therefore also within the plaque. The weird smell is likely due to the contaminant.

Hope this helps.

Vic

Thiel
Hello,
I think that we are having a similar problem to the post above that says "D29 plaques barely visible on M. smeg" but I'm not sure.
We are doing the phage isolation for the 2nd semester (so I'm not very experienced) and the students were on their 2nd round of purifications and starting to do the 3rd round and then make webbed plates. Our host is M. foliorum. I take new M. foliorum out of the -80C every 2 weeks, streak a plate and then shake it to make new stocks for them to use. The entire class started getting very similar looking plaques, that were difficult to see and smelled weird. (See image). The plates labeled "old" bacteria is what we had been using. The plates labeled "new" bacteria was when we picked a new colony from the same streak plate and it starting giving us the odd results. The difference in the incubation times was because we thought if we gave the 10^-3 plate a little more time- it would fully lyse, but it didn't.

We picked a another new colony from the M. foliorum streak plate, grew it up and it seems to have fixed the problem (no pic). Just in case, I made a new M. foliorum streak plate from a different glycerol stock and will be growing that up for future use for the students.

So we might have fixed the problem, but what do you think happened? Did we get lucky and pick a mutant colony? Any insight would be awesome. I'll let you know if the problem is not fixed.

Posted in: Phage Discovery/Isolation → Phages suddenly not fully lysing on M. foliorum

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