SEA-PHAGES | All posts created by viknesh

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

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

Link to this post \| posted 13 Sep, 2022 23:38
viknesh	cellokiwi So when you pass it, it goes back to yellow for a bit then turns orange again. We sadly don't have the resources to check 16S. Is there a resource for this through the program? The first time we saw it was in a lysogen so I just attributed it to wherever the genome had integrated itself messing with the color. Now, there has been zero change whatsoever. Same media, same CaCl2, same buffer, same incubator, same everything. One colleague suggested it is a carotenoid response to oxidative stress? Does that sound possible? Since we're using the same everything what would suddenly be stressing them out that wasn't before? In the picture the one on the left is a normal Arthrobacter culture and the one on the right is just starting to turn orange. They get darker than that. Hi Alison, Since we began using M . foliorum for phage-hunting in the SEA, I've yet to hear from anyone about it turning orange. Can you confirm the following? 1. If you streak a plate from your freezer/glycerol stock and incubate the plate at 30C, colonies start out yellow but then turn orange? How long before they turn yellow, and how long before they turn orange? If you have photos, please share. 2. If you setup a culture, they saturated culture is yellow but then eventually turns orange? If so, as before, please share the timing of these color changes. 3. If you streak a plate from the culture (from #2), colony formation and timin gof color changes proceed similar to streaking from the glycerol? 4. You have several phages that can still form plaques when plated with bacteria from the orange culture? Thanks. Vic Edited 13 Sep, 2022 23:39

Link to this post | posted 13 Sep, 2022 23:38

viknesh

cellokiwi
So when you pass it, it goes back to yellow for a bit then turns orange again. We sadly don't have the resources to check 16S. Is there a resource for this through the program? The first time we saw it was in a lysogen so I just attributed it to wherever the genome had integrated itself messing with the color. Now, there has been zero change whatsoever. Same media, same CaCl2, same buffer, same incubator, same everything. One colleague suggested it is a carotenoid response to oxidative stress? Does that sound possible? Since we're using the same everything what would suddenly be stressing them out that wasn't before?

In the picture the one on the left is a normal Arthrobacter culture and the one on the right is just starting to turn orange. They get darker than that.

Hi Alison,

Since we began using M . foliorum for phage-hunting in the SEA, I've yet to hear from anyone about it turning orange. Can you confirm the following?

1. If you streak a plate from your freezer/glycerol stock and incubate the plate at 30C, colonies start out yellow but then turn orange? How long before they turn yellow, and how long before they turn orange? If you have photos, please share.

2. If you setup a culture, they saturated culture is yellow but then eventually turns orange? If so, as before, please share the timing of these color changes.

3. If you streak a plate from the culture (from #2), colony formation and timin gof color changes proceed similar to streaking from the glycerol?

4. You have several phages that can still form plaques when plated with bacteria from the orange culture?

Thanks.
Vic

Edited 13 Sep, 2022 23:39

Posted in: Arthrobacter → Orange Cells

Link to this post \| posted 23 Aug, 2022 16:20
viknesh	A for effort and yAy for a cluster that with congruent calls! debbie Hi all, As of today, June 17, 2022, there are 88 Cluster EE genomes in our records. They are closely related genomes that contain only ~28 genes. In an effort to make the records congruent, my students and I have reviewed ~75 of them and revised 71 of the records. The template we used is attached here, along with a list of 71 records that we touched. We spent an abundant amount of time on the helix-turn-helix DNA binding proteins at the right end of the genome. We investigated 'types' of helix-turn-helix choices and decided the best path is to call them helix-turn-helix DNA binding proteins. August 23, 2022 - I just received word that the genomes that we sent to GenBank have been processed. This cluster just might have congruent calls! Thanks Vic! debbie Edited 23 Aug, 2022 18:40

Link to this post | posted 23 Aug, 2022 16:20

viknesh

A for effort and yAy for a cluster that with congruent calls!

debbie
Hi all,
As of today, June 17, 2022, there are 88 Cluster EE genomes in our records. They are closely related genomes that contain only ~28 genes. In an effort to make the records congruent, my students and I have reviewed ~75 of them and revised 71 of the records. The template we used is attached here, along with a list of 71 records that we touched. We spent an abundant amount of time on the helix-turn-helix DNA binding proteins at the right end of the genome. We investigated 'types' of helix-turn-helix choices and decided the best path is to call them helix-turn-helix DNA binding proteins.
August 23, 2022 - I just received word that the genomes that we sent to GenBank have been processed. This cluster just might have congruent calls!

Thanks Vic!
debbie

Edited 23 Aug, 2022 18:40

Posted in: Cluster EE Annotation Tips → Genome Curation - a must read!

Link to this post \| posted 05 May, 2022 17:50
viknesh	bgibb There used to be three protocols for setting up TEM samples, but now there is only one 8.1c (parafilm drop method) in the Discovery guide and Instructor guide. Is there a reason that the other protocols were removed? I've had good luck with the pelco tab method. Bryan, I see all 3 protocols in the Discovery Guide - https://seaphagesphagediscoveryguide.helpdocsonline.com/8-0-toc

Posted in: Phage Discovery/Isolation → Electron microscopy protocols

Link to this post \| posted 25 Jan, 2022 15:50
viknesh	sahas Hi I was wondering if anyone can recommend a place/lab to send our samples for electron microscopy? We haven't been successful in finding a resource yet. Any help will be highly appreciated. Thanks Sangha Similar to Cathy's suggestion, you could also reachout to UMBC's Imaging Facility. If you would like to cehck on their pricing and availability, you can email their facility director, Tagide deCarvalho at tagided@umbc.edu. Please copy me on that email to Tagide.

Link to this post | posted 25 Jan, 2022 15:50

viknesh

sahas
Hi
I was wondering if anyone can recommend a place/lab to send our samples for electron microscopy? We haven't been successful in finding a resource yet.
Any help will be highly appreciated.
Thanks
Sangha

Similar to Cathy's suggestion, you could also reachout to UMBC's Imaging Facility. If you would like to cehck on their pricing and availability, you can email their facility director, Tagide deCarvalho at tagided@umbc.edu. Please copy me on that email to Tagide.

Posted in: General Message Board → Electron Microscopy

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