SEA-PHAGES | All posts created by viknesh

Link to this post \| posted 24 Feb, 2024 15:26
viknesh	jdaft@leeuniversity.edu Has anyone ever had an issue with thermophile contamination in their top agar? We've cultured the thermophile to a PYCa plate and growth only occurs at 60C and not 37C. We think we narrowed it down to our calcium chloride and/or dextrose. Both were remade and filtered and we still are having contamination issues. Our host bacteria is M. foliorum, so our thermophile doesn't appear to be interfering with it's growth, but we still don't like have another bacteria in there with our host culture. ~Joe Can't say I've heard from the community about an issue like this. Can you share more about how you discovered it, since it doesnt grow at 37? Photos would be great too. Vic

Link to this post | posted 24 Feb, 2024 15:26

jdaft@leeuniversity.edu
Has anyone ever had an issue with thermophile contamination in their top agar? We've cultured the thermophile to a PYCa plate and growth only occurs at 60C and not 37C. We think we narrowed it down to our calcium chloride and/or dextrose. Both were remade and filtered and we still are having contamination issues. Our host bacteria is M. foliorum, so our thermophile doesn't appear to be interfering with it's growth, but we still don't like have another bacteria in there with our host culture.
~Joe

Can't say I've heard from the community about an issue like this. Can you share more about how you discovered it, since it doesnt grow at 37? Photos would be great too.

Vic

Posted in: Phage Discovery/Isolation → Top Agar contamination

Link to this post \| posted 11 Jan, 2024 23:31
viknesh	Below is a message from David Asai to the SEA community I want to take a moment to share with you some personal news. I am retiring from HHMI today. For more than 15 years, it has been my privilege to be part of an amazing team committed to improving science education. I am pleased that the good work continues with renewed focus as part of the HHMI Center for the Advancement of Science Leadership and Culture (www.hhmi.org/equitable-science). HHMI’s Science Education Alliance has the immodest goal of revolutionizing the way science is learned. We look to a future when all students – regardless of where they come from and where they are going – learn science in an environment in which they feel that they belong and provides them the resources to be successful. You have much of which to be proud. When the SEA was launched in 2008, we were 12 colleges and universities. Today, there are more than 160 active member schools where 5,500+ students and 600+ faculty annually engage in discovery-based learning. More impressive and important than the numbers is that SEA students from all backgrounds and all institution types score significantly higher than students in traditional laboratory courses in the indicators of belonging and persistence in science. There are two keys to our success, the science and you. We prioritize the high quality of the science, and you commit to being part of an active community of practice devoted to student learning. My sincere gratitude to all of you for your deep commitment and hard work creating new learning opportunities for our students. Thank you to the SEA team – Billy Biederman, Kaylia Edwards, Danielle Heller, Pushpa Ramakrishna, Vic Sivanathan, and Bethany Wise. I am eager to see what you all accomplish together. Happy new year and Aloha. David Asai david.asai@outlook.com Edited 11 Jan, 2024 23:32

Link to this post | posted 11 Jan, 2024 23:31

viknesh

Below is a message from David Asai to the SEA community

I want to take a moment to share with you some personal news. I am retiring from HHMI today. For more than 15 years, it has been my privilege to be part of an amazing team committed to improving science education. I am pleased that the good work continues with renewed focus as part of the HHMI Center for the Advancement of Science Leadership and Culture (www.hhmi.org/equitable-science). HHMI’s Science Education Alliance has the immodest goal of revolutionizing the way science is learned. We look to a future when all students – regardless of where they come from and where they are going – learn science in an environment in which they feel that they belong and provides them the resources to be successful. You have much of which to be proud. When the SEA was launched in 2008, we were 12 colleges and universities. Today, there are more than 160 active member schools where 5,500+ students and 600+ faculty annually engage in discovery-based learning. More impressive and important than the numbers is that SEA students from all backgrounds and all institution types score significantly higher than students in traditional laboratory courses in the indicators of belonging and persistence in science. There are two keys to our success, the science and you. We prioritize the high quality of the science, and you commit to being part of an active community of practice devoted to student learning. My sincere gratitude to all of you for your deep commitment and hard work creating new learning opportunities for our students. Thank you to the SEA team – Billy Biederman, Kaylia Edwards, Danielle Heller, Pushpa Ramakrishna, Vic Sivanathan, and Bethany Wise. I am eager to see what you all accomplish together.

Happy new year and Aloha. David Asai david.asai@outlook.com

Edited 11 Jan, 2024 23:32

Posted in: General Message Board → A Message from David Asai

Link to this post \| posted 05 Dec, 2023 18:51
viknesh	RyanWheeler Hello everyone, I am looking purchase materials for the DNA extraction and I am a little stuck on the item "nuclease mix". Does this refer to a mix of DNase and RNase or some specific product? Which brand of either works the best in your experience? Thanks, Ryan Hi Ryan, The nuclease mix contains both DNaseI and RNaseA – see the recipe card in the Phage Discovery Guide, which has a recipe for every reagent that needs to be prepared. The recipe for the nuclease mix does have you prepare DNaseI and RNaseA separately, using the manufacturer's guidelines, before combining them to prepare the mix. Both the ordering guide and Fisher Quote lists an example of DNaseI and RNaseA for purchase. We're open to trying nucleases from other sources too, especially if they are cheaper. Edited 05 Dec, 2023 18:52

Link to this post | posted 05 Dec, 2023 18:51

viknesh

RyanWheeler
Hello everyone,
I am looking purchase materials for the DNA extraction and I am a little stuck on the item "nuclease mix". Does this refer to a mix of DNase and RNase or some specific product? Which brand of either works the best in your experience?
Thanks,
Ryan

Hi Ryan,

The nuclease mix contains both DNaseI and RNaseA – see the recipe card in the Phage Discovery Guide, which has a recipe for every reagent that needs to be prepared. The recipe for the nuclease mix does have you prepare DNaseI and RNaseA separately, using the manufacturer's guidelines, before combining them to prepare the mix. Both the ordering guide and Fisher Quote lists an example of DNaseI and RNaseA for purchase.

We're open to trying nucleases from other sources too, especially if they are cheaper.

Edited 05 Dec, 2023 18:52

Posted in: Phage Discovery/Isolation → Best nuclease mix?

Link to this post \| posted 05 Oct, 2023 15:42
viknesh	engstrom Hi Community, We have been stopped in our tracks here at Monmouth, by a very strange issue. Our PYCa top-agar is useless as our CaCl2 is consistently precipitating. We made a new 1M stock of course. We made new dextrose as well. We played with our autoclaving times. And we are not having any trouble with plate agar. At a loss. My question–can we use top-agar for plaque assays if we omit the CaCl2? This would seem to be the quickest route to solving our problem and we will make a test of it soon. Other suggestions? Thanks! Eric, thanks for sharing. We too were stumped this year with calcium precipitating in our PYCa top agar. It is incredibly frustrating – so sorry you too are experiencing this. I believe I heard from one other person about this too. I wonder if it has to do with more recent batches of media components. I'm not sure what the best way forward is, but one thing we've found that minimizes the likelihood of precipitation is to first autoclave the media without calcium chloride (and we all typically do), then allow it too cool all the way do about 60C (before the top agar begins to set), then add the calcium chloride (and in our case, dextrose too), swirl, and place it in the water bath. Allowing it to solidify and melting it in the microwave seems to promote the precipitation. Out of 20 x 100 ml bottles of top agar prepared as a single batch, all at once, we find that a majority will not crash for at least 2 - 3 days. We have noticed that in those that do not crash, we see flaky crystals forming, presumably calcium precipitating in a more ordered fashion. The randomness of this phenomenon across the aliquots suggests that we're at some tipping point, and that very slight variations can trigger precipitation. I suspect pH can have done nothing to verify this. We'll see what we can learn. in the meanwhile, good luck and keep us up to date. Thanks. Edited 05 Oct, 2023 15:46

Link to this post | posted 05 Oct, 2023 15:42

viknesh

engstrom
Hi Community,

We have been stopped in our tracks here at Monmouth, by a very strange issue. Our PYCa top-agar is useless as our CaCl2 is consistently precipitating. We made a new 1M stock of course. We made new dextrose as well. We played with our autoclaving times. And we are not having any trouble with plate agar. At a loss. My question–can we use top-agar for plaque assays if we omit the CaCl2? This would seem to be the quickest route to solving our problem and we will make a test of it soon. Other suggestions?

Thanks!

Eric, thanks for sharing. We too were stumped this year with calcium precipitating in our PYCa top agar. It is incredibly frustrating – so sorry you too are experiencing this. I believe I heard from one other person about this too. I wonder if it has to do with more recent batches of media components. I'm not sure what the best way forward is, but one thing we've found that minimizes the likelihood of precipitation is to first autoclave the media without calcium chloride (and we all typically do), then allow it too cool all the way do about 60C (before the top agar begins to set), then add the calcium chloride (and in our case, dextrose too), swirl, and place it in the water bath. Allowing it to solidify and melting it in the microwave seems to promote the precipitation. Out of 20 x 100 ml bottles of top agar prepared as a single batch, all at once, we find that a majority will not crash for at least 2 - 3 days. We have noticed that in those that do not crash, we see flaky crystals forming, presumably calcium precipitating in a more ordered fashion. The randomness of this phenomenon across the aliquots suggests that we're at some tipping point, and that very slight variations can trigger precipitation. I suspect pH can have done nothing to verify this. We'll see what we can learn. in the meanwhile, good luck and keep us up to date. Thanks.

Edited 05 Oct, 2023 15:46

Posted in: Phage Discovery/Isolation → Precipitation of CaCl2 in PYCA top-agar

Link to this post \| posted 06 Apr, 2023 17:27
viknesh	Thiel Hi, I can share a photo but Kristen shared a great suggestion with me that worked. We put 150uL of phage buffer in a tube and then picked 15-20 plaques from the same plate. We had done 4 purifications so we were confident that it was pure. We put the 15-20 plaques in the 150uL of phage buffer. We then did serial dilutions, infected the bacteria and plated as normal the 10^0 through 10^-4 and the 10^-3 was very nicely webbed. Great!

Link to this post | posted 06 Apr, 2023 17:27

viknesh

Thiel
Hi, I can share a photo but Kristen shared a great suggestion with me that worked. We put 150uL of phage buffer in a tube and then picked 15-20 plaques from the same plate. We had done 4 purifications so we were confident that it was pure. We put the 15-20 plaques in the 150uL of phage buffer. We then did serial dilutions, infected the bacteria and plated as normal the 10^0 through 10^-4 and the 10^-3 was very nicely webbed.

Great!

Posted in: Phage Discovery/Isolation → Low plate titer issues

Link to this post \| posted 31 Mar, 2023 14:37
viknesh	Thiel I've had this same issue with a new turbid plaque, PhriedEgg. I'm not sure of an answer but if anyone has suggestions then that would be great. Thanks. Hi Sarah, Would you be able to share photos of what you are seeing? Vic

Posted in: Phage Discovery/Isolation → Low plate titer issues

Link to this post \| posted 05 Dec, 2022 05:17
viknesh	stpage Vic wrote: "Like preparing webbed plates, it is important that not all the host bacteria is lysed early. It is therefore important that you setup the experiment early in the day and check on the culture before you leave in the evening. If the culture is clear (and the control is cloudy), then you've run out of host bacteria for phage amplification." Hi, what is the concern with "early lysis"? Is it just that it limits phage amplification or that too many bacterial degradative products are released early or…..? (just wondering, we've had trouble getting students to be consistent with timing) Yup - if all host cells are lysed after only a few rounds of infection, the amplification will be low. You really need to get to later rounds of infection… 7 ish and above in order to have a high titer.

Link to this post | posted 05 Dec, 2022 05:17

viknesh

stpage
Vic wrote: "Like preparing webbed plates, it is important that not all the host bacteria is lysed early. It is therefore important that you setup the experiment early in the day and check on the culture before you leave in the evening. If the culture is clear (and the control is cloudy), then you've run out of host bacteria for phage amplification."

Hi, what is the concern with "early lysis"? Is it just that it limits phage amplification or that too many bacterial degradative products are released early or…..? (just wondering, we've had trouble getting students to be consistent with timing)

Yup - if all host cells are lysed after only a few rounds of infection, the amplification will be low. You really need to get to later rounds of infection… 7 ish and above in order to have a high titer.

Posted in: Phage Discovery/Isolation → Protocol to grow phage in culture?

Link to this post \| posted 10 Nov, 2022 16:33
viknesh	plconnerly Vic - we're using all samples with a titer of at least 1 x 10^9. Typically, we can get the minimum 40 ng/ul from a lysate at 5x10^9 pfu/ml, though of course that will vary from phage to phage. Including a ZnCl2 precipitation step has worked well for many, with University of Ottawa now including this as a standard step for all their DNA extractions. It does take a little getting used to, and it is very important to be resuspending the pellet very quickly after the spin, and in EDTA. The precipitation step is harsh and phage DNA is rapidly released after the spin, making them accessible to the nuclease before the denaturant is added. I havent tried this, but perhaps it makes sense to add EDTA to the nuclease-treated lysate before adding ZnCl2. As you troubleshoot, I'd recommend prepping DNA from one lysate both ways (with and without ZnCl2), side by side, to see if the precipitation step works for you. Let me know if you want to chat via Zoom.

Link to this post | posted 10 Nov, 2022 16:33

viknesh

plconnerly
Vic - we're using all samples with a titer of at least 1 x 10^9.

Typically, we can get the minimum 40 ng/ul from a lysate at 5x10^9 pfu/ml, though of course that will vary from phage to phage. Including a ZnCl2 precipitation step has worked well for many, with University of Ottawa now including this as a standard step for all their DNA extractions. It does take a little getting used to, and it is very important to be resuspending the pellet very quickly after the spin, and in EDTA. The precipitation step is harsh and phage DNA is rapidly released after the spin, making them accessible to the nuclease before the denaturant is added. I havent tried this, but perhaps it makes sense to add EDTA to the nuclease-treated lysate before adding ZnCl2.

As you troubleshoot, I'd recommend prepping DNA from one lysate both ways (with and without ZnCl2), side by side, to see if the precipitation step works for you.

Let me know if you want to chat via Zoom.

Posted in: Phage Discovery/Isolation → DNA Extraction Troubleshooting - Can we skip the nuclease treatment?

Link to this post \| posted 09 Nov, 2022 19:46
viknesh	plconnerly We could use any and all advice about DNA extraction from Gordonia rubripertincta phages. Last year we tried and troubleshot the Wizard DNA Cleanup kit column method with no success. We did phenol:chloroform:isoamyl alcohol as a last resort and managed to get useable DNA from 1 phage. This year we're trying the ZnCl2 method and initial runs have not worked. We've got some troubleshooting planed and one idea we have is to leave out the initial nuclease step. Is that allowed? Have other folks tried that? Can DNA extracted without nuclease treatment be used for sequencing? Thanks! Pam - IU Southeast I'm hoping Dan Russell will chime in here but I believe it is very important to include the nucelase treatment step. Otherwise, the bacterial DNA to phage DNA ratio might be too high, resulting in insufficient sequencing depth for the phage genome. Can I ask about the titer of the samples that are resulting is low DNA extraction?

Link to this post | posted 09 Nov, 2022 19:46

viknesh

plconnerly
We could use any and all advice about DNA extraction from Gordonia rubripertincta phages. Last year we tried and troubleshot the Wizard DNA Cleanup kit column method with no success. We did phenol:chloroform:isoamyl alcohol as a last resort and managed to get useable DNA from 1 phage. This year we're trying the ZnCl2 method and initial runs have not worked. We've got some troubleshooting planed and one idea we have is to leave out the initial nuclease step. Is that allowed? Have other folks tried that? Can DNA extracted without nuclease treatment be used for sequencing?
Thanks!
Pam - IU Southeast

I'm hoping Dan Russell will chime in here but I believe it is very important to include the nucelase treatment step. Otherwise, the bacterial DNA to phage DNA ratio might be too high, resulting in insufficient sequencing depth for the phage genome.

Can I ask about the titer of the samples that are resulting is low DNA extraction?

Posted in: Phage Discovery/Isolation → DNA Extraction Troubleshooting - Can we skip the nuclease treatment?

Link to this post \| posted 21 Sep, 2022 17:09
viknesh	edoddmoh@uottawa.ca Hi! Is there is a reason the smeg top agar is prepared at 2X, and then diluted to 1X? I'm wondering if I can prepare it as a 1L volume, autoclave, and then aliquot into smaller bottles (as with the PYCa top agar). We are phage-hunting with smeg for the first time this semester. Thanks! Liz Hi Liz, Part of the reason top agar is prepared at 2x is so that experiemnts that require the use or large samples of phage (in buffer) do not overly dilute the nutrient in the medium. For our general PHAGES protocols, you should be fine preparing top agar at 1x. 7H9 medium has lots of salts in it, and calcium chloride tends to precipitate out over time. For this reason, calcium chloride is only added before use. Even if you prepare 1x top agar, you might want to only add calcium chloride before use. If you do add it early, please let us know how long it takes for the precipitates to form so that we can share with others. Vic

Link to this post | posted 21 Sep, 2022 17:09

viknesh

edoddmoh@uottawa.ca
Hi! Is there is a reason the smeg top agar is prepared at 2X, and then diluted to 1X? I'm wondering if I can prepare it as a 1L volume, autoclave, and then aliquot into smaller bottles (as with the PYCa top agar). We are phage-hunting with smeg for the first time this semester. Thanks!
Liz

Hi Liz,

Part of the reason top agar is prepared at 2x is so that experiemnts that require the use or large samples of phage (in buffer) do not overly dilute the nutrient in the medium. For our general PHAGES protocols, you should be fine preparing top agar at 1x. 7H9 medium has lots of salts in it, and calcium chloride tends to precipitate out over time. For this reason, calcium chloride is only added before use. Even if you prepare 1x top agar, you might want to only add calcium chloride before use. If you do add it early, please let us know how long it takes for the precipitates to form so that we can share with others.

Vic

Posted in: Phage Discovery/Isolation → 2X smeg top agar

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