SEA-PHAGES | All posts created by viknesh

Link to this post \| posted 03 Apr, 2024 12:04
viknesh	Applications for BioInteractive’s Ambassador Academy are now open! We’re looking for educators with diverse backgrounds and teaching contexts to apply to this multi-year professional development program to join our Ambassador Community. Applications consist of two sections: completion of eligibility questions and completion of an application package. To apply, educators must complete the eligibility questions by May 7, 2024. The remainder of the application package is due by September 12, 2024. More about the Academy, including eligibility criteria, benefits of participation, and a link to the application, can be found on the Academy webpage. Edited 03 Apr, 2024 12:06

Link to this post | posted 03 Apr, 2024 12:04

Applications for BioInteractive’s Ambassador Academy are now open! We’re looking for educators with diverse backgrounds and teaching contexts to apply to this multi-year professional development program to join our Ambassador Community.

Applications consist of two sections: completion of eligibility questions and completion of an application package. To apply, educators must complete the eligibility questions by May 7, 2024. The remainder of the application package is due by September 12, 2024.

More about the Academy, including eligibility criteria, benefits of participation, and a link to the application, can be found on the Academy webpage.

Edited 03 Apr, 2024 12:06

Posted in: General Message Board → BioInteractive Ambassador Eligiblity is Open until May 7, 2024. Apply Now

Link to this post \| posted 31 Mar, 2024 02:59
viknesh	tperezmo Hi all, I recently switched one section to work on A. atrocyaneus while the others work on A. globiformis. For A. globiformis, we always have really good DNA yields 40 - 100 ng per ul. However, for A. atrocyaneus, we are not hitting even 20 ng / ul. I have used the original protocol with nucleases, SDS, EDTA, Proteinase K. The results are for all students in the class, for 2 semesters now. When trying to do the restriction digests, we can barely see a band in the controls. I have looked at the Microbacterium forum but there it seems it's a lysate concentrations. All my students have at least 1 x 10^8 or more. Any thoughts? Thank you! Hi Tiara, I don't have experience working with A. atrocyaneus, but a couple of things. 1. Have you considered using ZnCl2 precicptation? It seems to do the trick for folks with low yield. Also, if your titers are generally around 10^8, you might want to see if your titer is the issue. Do you still get low DNA yield from a lysate with a titer of 5 x 10^9 pfu/ml? 2. A quick search on phagesDB shows that Pitt and UCLA have submitted DNA for a few phages isolated on this host last year. I'd recommend pinging SEA faculty Kristen Butela and Amanda Friese for input (and have the respond on the forum so we capture it here for others too). Thanks. Vic

Link to this post | posted 31 Mar, 2024 02:59

viknesh

tperezmo
Hi all,

I recently switched one section to work on A. atrocyaneus while the others work on A. globiformis. For A. globiformis, we always have really good DNA yields 40 - 100 ng per ul. However, for A. atrocyaneus, we are not hitting even 20 ng / ul. I have used the original protocol with nucleases, SDS, EDTA, Proteinase K. The results are for all students in the class, for 2 semesters now.
When trying to do the restriction digests, we can barely see a band in the controls.
I have looked at the Microbacterium forum but there it seems it's a lysate concentrations. All my students have at least 1 x 10^8 or more.
Any thoughts? Thank you!

Hi Tiara,

I don't have experience working with A. atrocyaneus, but a couple of things.

1. Have you considered using ZnCl2 precicptation? It seems to do the trick for folks with low yield. Also, if your titers are generally around 10^8, you might want to see if your titer is the issue. Do you still get low DNA yield from a lysate with a titer of 5 x 10^9 pfu/ml?

2. A quick search on phagesDB shows that Pitt and UCLA have submitted DNA for a few phages isolated on this host last year. I'd recommend pinging SEA faculty Kristen Butela and Amanda Friese for input (and have the respond on the forum so we capture it here for others too).

Thanks.
Vic

Posted in: Phage Discovery/Isolation → A atrocyaneus DNA extraction

Link to this post \| posted 18 Mar, 2024 14:53
viknesh	Thanks for sharing, Joe. I think this is going to be ver helpful for other faculty if they are seeing all their top agar get contaminated on a regular basis. Hvae you been able to narrow it down to your dextrose or calcium chrloride solution. Any more info you can share will be helpful for the community.

Posted in: Phage Discovery/Isolation → Top Agar contamination

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

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

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?

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