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Invisiblec10h12n2o
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Registered: 01/21/15
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c10's Agar Guide + Tips & Tricks (post #1000) * 43
    #24330957 - 05/18/17 09:00 PM (3 years, 4 months ago)

Disclaimer: i am a certifiable agar addict. lots of people ask me for the specifics of how i do agar, and rather than repeating myself every day, here is a convenient guide to the way i do agar, as well as some tips and tricks i have picked up. This is simply my post #1000 way of sharing what i have learned over 20k+ plates, and is not meant to be a comprehensive agar guide, bodhi's comprehensive agar tek does a fantastic job of that, and should be required reading for anyone starting agar. lots of what i do is less than ideal; i will try to point out where i am cutting corners, but i welcome any criticism, questions, additions, refutations, or discussion you have to add :smile:

c10's Agar Guide + Tips & Tricks

or M.A.G.A. : Make Agar Great Again!



couple weeks' supply of plates. make that mailman work for his money.


INTRO:
Agar is ubiquitous in professional mycology (or any form of microbiology for that matter). In the old days, when people were just figuring out the bulk teks, pros had been using agar for decades. For this reason, many people have been intimidated by agar and its prominent use on the cutting edge of mycology, thinking it was a tool reserved for people with proper labs and degrees, far beyond the scope of the average hobbyist. this was very misguided, because in reality making agar is no more difficult than making jello shots, and there are lots of agar techniques that will benefit even the newest of novices. agar removes all of the guess-work and virtually guarantees success. for this reason, these days we like to advise people to START with agar as soon as they enter the hobby

Quote:

Niccolo Machiavelli said:
The wise man does at once what the fool does finally.




do not be afraid of failing. agar gives us the chance to do culture work in very small spaces and run experiments with lots of data points. plates are CHEAP, so dont get attached to them. dont get discouraged by contamination. in fact, you can learn FAR more from failure than you can from success. if you think critically, you can learn a LOT from a contaminated plate: what techniques DONT work, what you are doing wrong, how contams behave on agar, how myc reacts to these stimuli, how extreme your clean technique needs to be, etc.. by comparison, ESPECIALLY with a species as hardy as cubensis, success can teach us relatively little. so learn to see your failures as opportunities to learn: make the most out of them. stay with it and you will be consistently successful, just be patient and persistent while keeping an open-minded 

Regarding SABs and FHs:


To do agar work, you are going to need to be somewhat familiar with clean technique, etc., and will need either a SAB (still air box) or a FH (laminar flow hood/cabinet). whichever route you go, its important to understand the basic principles of how the device works.

SABs:
For a SAB, the principle at work is STILL AIR, meaning you can NEVER get the SAB clean enough to do sterile work (it could be quite dirty and function just fine). STILL AIR (as opposed to STERILE BOX) is why a SAB works. in a SAB we are counting on gravity to pull particles/contams out of the air, and the SAB blocks interference/turbulence from outside.

For these reasons, it is crucial that you understand that contams are concentrated on the bottom of a SAB, and for this reason i STRONGLY recommend placing a damp towel in the bottom of the SAB and working on a cookie rack 2-3" above it. bodhi accomplishes the same thing by using some petris as spacers to raise his work surface off the horizontal floor surface. the idea is put some space between the entry to your clean culture dish (mouth of your petri) and the horizontal surface full of contams, and the towel helps trap contams and keep them from being stirred up by the movements of your hands.

this is also one of the reasons some people have better luck with no-pour/pastyplates than standard petris, since pp5 containers ALWAYS have a MUCH higher rim than petris (usually a matter of multiple inches) and thus have a wider space between the opening to the agar dish and the dirty floor. pouring petris in a SAB with no space between the dish you are workin in and the floor can easily go badly. that space can make a HUGE difference. over the course of about 5k plates without a towel+rack and another 15k with them, the single change of adding the towel+rack brought my success rate from around 94% to 99% clean (from 1/20 to 1/100)

if you need instructions for making a SAB,
check out bodhi's simple AF SAB tek

these are two SABs i built before i constructed my FH, a double hatch one (missing the towel, i hadnt learned that yet and was using contact mats to avoid slippage), and a totally custom one i built in this thread out of plexiglass i found on the side of the road :lol:

2 types of custom SABs i built


that custom one actually turned out really great, i LOVED that thing, it was built into a closet to leverage the walls for extra blocking of air currents and to be easily concealable, and it was plenty tall for bags :smile: . but looking back, i would advise going the simple AF route, then later building a custom SAB or FH to your needs if you still want one (basic SAB is all you need)

Flow Hoods/Cabinets:
in a laminar flow hood the air is passed through a HEPA (High Efficiency Particulates Air) filter which removes all airborne contamination to maintain sterile conditions. A laminar flow hood consists of a prefilter, a blower, and a HEPA filter.

so this is TOTALLY different from the SAB in terms of the principles being leveraged to allow for sterile work. in a SAB we rely on STILL AIR, but in a FH (flow hood/cabinet) we rely on a laminar column of sterile air. basically, you dont want to let anything potentially dirty get in between the sterile airflow and your clean work (like a petri dish)

my beloved flow hood


here are a few FH links for anyone wanting to build one:
c10s flow hood build
stonesun's FH tek
mushpunx FH build

i have REALLY been enjoying mine, it has been a joy to work with. i have literally only seen 1 plate contam since i built it (out of over a thousand) and it was one i dropped on the floor (outside the hood):lol:

Agar Prep:


here is how i prep agar, a liter at a time. i do a few things that arent exactly kosher, and will point those out.

getting materials together


like everything else we do in this hobby, it is CRUCIAL that we understand the point of what we are doing. when working with agar, we want a slightly nutritious medium which supports controlled growth in 2 dimensions. for this reason, nearly any kind of media the myc can digest can work fine (historically people have used cardboard and all kinds of stuff). These days, most people prefer either MEA (the standard in pro mycology), PDA (potato dextrose agar), or GSA (grain soak agar), but since we are using them to accomplish the same thing it doesnt really matter which recipe you go with as long as you have the right balance of agar-agar and sugar.

many of us like to use "half strength" agar with half the normal sugar content, to encourage myc to spread out and look for food rather than grubbing out and staying put, and make it easier for us to spot sectoring and other differences in mycelium characteristics. my preferred recipe is a "half strength" recipe of MEA (20g agar-agar, 15g light malt extract, 1L h2o), which i sometimes alternate with PDA or GSA.

i have experimented a LOT with different agar recipes and preparations, and found the simplest recipes to be the best for uniform results. peptone and yeast would often make the myc act weird, and eventually i quit supplementing with things like that because i realized that focusing on the nutrition of the agar (beyond what is required for healthy, consistent growth) kinda misses the point.

if you plan to be doing lots of agar work, and want to be able to draw meaningful conclusions about how one culture looks/acts vs another one, it is important that you standardize your agar recipe and prep (find something you like and stick to it), otherwise you wont be able to know whether you are observing a characteristic of the culture or a difference due to a different agar recipe

antibiotics are totally not necessary in this hobby, since you can clean up pretty much anything with well-timed transfers. i do keep gentamicin and chloramphenicol on hand, and occasionally use it, but it should absolutely not be a matter of course. the only thing i use it for is the final dish before making long-term master slants, to knock out any hidden bacteria before we put it into storage (just in case). i do not use antibiotics in the master slants, just the final plate before it, and usually dont use them even in these cases (they arent necessary). i have also used it before doing a2g (in the rare occasions when im trying to troubleshoot suspected hidden bacteria), or when a print is so incredibly dirty that the bacteria suppresses the myc from growing at all on standard agar (ive only seen this twice, and i think someone wiped their ass with the print).

i have ordered agar-agar from lots of suppliers, and dont see much difference in the bacteriological (EXPENSIVE) kind, the health food store kilo kind, and the telephone brand kind. as long as it is pure agar-agar, white/offwhite powder, it should be fine. i prefer to buy in bulk, but most often have ended up using telephone brand from the asian stores

old faithful


To make my standard MEA, i like to use a 2L beaker to mix everything in. First, i tare (zero) a food scale with the beaker on it, then weigh out 20g (ish, a little more/less wont hurt) of agar-agar like this:

weighing agar inside mixing beaker


next, i tare the scale again and weigh out 15g (ish) of light malt extract and mix the 2 dry powders together:

weighing malt extract inside mixing beaker


then add a small amount of cold water (tap is fine), just enough to mix the powders into a thick liquid. i like to use a glass stir rod with a rubber policeman on the end (from a lab supply store) to mix this suspension as evenly as possible:

concentrated agar suspension


and then add cold water until the total volume is 1 liter and stir well:

1L of agar suspension


now i add food coloring, which is usefult for 2 reasons: #1 it helps sort cultures visually for color coding, and #2 the coloration makes it much easier to spot healthy white mycelium and distinguish contaminants:

add food coloring (optional)


next i use a microwave to carefully boil the agar suspension and dissolve the mixture into a homogeneous solution. agar is VERY strange when it boils, it can easily start bubbling and boil over into a huge mess. for this reason, you need to keep a close eye on it and stir regularly. i like to get it to start boiling at full power, then reduce the microwave power down to 50% to maintain a controlled boil so it doesnt boil over. even at reduced power, i have to pop it open every few moments. i like to let it boil until i am sure everything has dissolved, then let it go a bit longer just to be sure

keep a close eye on this or you will have a huge mess


next, i take a towel and wrap it around the beaker so that i can handle it, and pour the contents through a funnel lined with cheesecloth, filling 2 1L media bottles halfway each (so they dont boil over). the purpose of the cheesecloth filter is to remove any clumps or foreign particles that might still be in your agar, to make sure everything is of an even consistency. this results in crystal clear agar that looks like glass, and makes pouring predictable. after filling, tighten the lids, then just barely crack them open so that steam can escape during the sterilization cycle

pouring agar through cheesecloth into media bottles, ready to sterilize


i like to sterilize my scalpel handles at the same time i do agar, usualy 8 or so at a time so that i dont have to do it often. i tear off a length of aluminum foil and fold it around a scalpel handle and fold it up into an envelope like shown. i then put a jar filled with these individually wrapped handles in the pressure cooker, with foil over the top so it doesnt fill up with water

individually wrapped scalpel handles for sterilization


these liter media bottles wont fit in my PC if i stand them straight up, so i have to get crafty to hold 2 of them. to accomplish this i stack 4 small jars against one edge and 2 jar rings along the other. i place the top side of the bottles onto the side elevated by the jars, and tilt the bottom edge against the rings, so that the media bottles do not come in contact with the edges of the pressure cooker

media bottles carefully loaded into pressure cooker


i then put hot water into the pressure cooker until it is close to level with the agar in the bottles, then close the lid and fasten it down. now we vent the PC by keeping the pressure valve open and turning the heat on until there is a constant plume of steam and reduce heat to medium-low to maintain this plume of steam. start a timer for 10 minutes and let it vent until the time is up


vent the PC for 10 minutes


when the 10 minutes is up, flip your valve (or apply your weight) and allow pressure to start building up. when it gets above 15 psi, start a timer for 20 minutes and maintain 15-18 psi for the duration of this time

PC for 20 minutes at 15-18 psi


WARNING: this is a little bit less than kosher, but i will provide the reasoning every guide says to let the PC zero out on its own. naturally the pressure falls once you kill the heat, and once it hits zero it goes below zero, then eventually back up. in my early work with agar, i had some contamination due to opening the PC when the pressure was negative, which led to sucking contams into my agar. what i like to do now is a little impatient, but i have not not seen any problems from it in the 15k to 20k plates i have done with this method. so here is what i like to do: i carefully watch the pressure as it falls, then right as it gets to zero, right before it crosses the line, i pop the vent. this means there is a small hiss of positive pressure being released from the valve (which is good, because it means nothing was sucked in), with less and less pressure as you get closer to zero, with negative pressure building if you wait too long (which sucks in contams). so right before zero, i put on protective gloves/mits, pop the valve, open the PC, and immediately tighten the lids on the media bottles and remove them

lids tightened and bottles removed from PC


now wait until it cools down to about 60 degrees Celsius. typically, i try to pour it before it starts gelling around 50 degrees. I like to use an infrared thermometer to check this, its really handy, but this is also the maximum where you can hold the bottle with your bare hands comfortably. so as if you dont have an IR thermometer, just wait until it has cooled down enough to pour comfortably

sterilized agar ready to pour


Pouring Plates:


now we move the agar to the our clean work area. im gonna be demonstrating with a FH, but same principles apply. you will notice, lots of people are successful even though they dont wear gloves, use a towel in their SAB, bathe (like ever lol), use iso, or whatever. this is because everyone's situation is different, and will have different requirements in terms of how extreme the sterile technique will need to be in a given environment. people dont necessarily have to do things perfectly to achieve success, and as you go you will learn which corners you can cut vs what is required in your environment. that said, when you are first working towards clean cultures/spawn, it is a good idea to be as clean as possible until you can reliably achieve clean spawn/cultures. so basically, when you see someone say they dont wear gloves, or dont use a towel, it doesnt mean you shouldnt (at least until you can establish that it doesnt matter in your case). its important to understand how these dynamics of sterile technique play out, and learn what works for you in your environment
 
work area for pouring plates


for a long time i would wear face masks, but my face always got sweaty/hot and uncomfortable after a while. eventually i started using a face shield, and i LOVE it. this thing is great for blocking your exhales (you can cough straight at the agar without worry) and also prevents hairs and skin cells from falling off your face. Another thing i like using is the black heavy duty gloves. these hold up a lot better than the standard gloves, especially over multiple uses. i reuse the same pair of gloves over and over, and wipe them down with alcohol before each session. i like to keep a rag soaked in 70% alcohol (this is dangerous so be careful) in a plastic bag in a container, so that the alcohol doesnt evaporate while im not using it. i pull this rag out and wipe things down with it regularly while working in the FH

face shield and alcohol rag wipe-down


now i pull out 60-80 plates worth of sleeves, and turn them upside down. using a retractable scalpel, i cut the end off each sleeve, then turn them right side up and pull the plastic sleeves off like this:

removing plates from sleeves


next i use the alcohol rag to wipe down my forearms and gloves, then carefully and thoroughly wipe the first media bottle. usually the lid is on very tight, so i use a strap wrench to loosen it, then i remove it by hand

strap wrench really helps break them open


then i center a stack of 20 plates and use the flat side of the bottle to line them up like so:

using the media bottle edge to line up the stack


next, i pick up the top 19 plates and the lid of the bottom one in my left hand, and pour agar into the bottom dish. if the plate needs to last more than long enough to transfer, i pour a layer that covers the whole floor of the dish, then replace the lid and move to the next plate (which is the kosher way to do it). usually, i try to maximize the number of plates i get per liter of agar, even at the expense of the agar possibly drying out in a few weeks (long after it has served its purpose). i do this by pouring about 1/3 to 1/2 the amount of agar required to cover the bottom of the dish, then i tilt the dish back and forth to make the agar cover the entire plate surface. this results in very thin agar medium which dries out easily, but also has some advantages (like not clogging up an LI needle with agar, since anything you inoculate with agar this thin will be almost pure myc, very little agar). picking up the plates is of course a possible contam vector, so i wouldnt recommend doing it this way until you can reliably achieve clean cultures (so that you will be able to tell if it causes problems). this is less risky in a SAB than a FH, since with a FH extra care needs to be taken to ensure that your hands dont get between the flow and your medium. another benefit of super thin plates is that transfers jump off onto receiving dishes faster, since there is less edge for them to colonize before it reaches the receiving dish. for this reason, even when using a standard thickness plate, i like to trim my transfers so that they are as thin as possible (cutting off the uncolonized agar underneath the myc) before transferring)

pouring plates super thin by using a tilting technique to spread the media. this is easiest when the media is relatively hot, because as it starts to gel and clumps form the surface will deform and wont be slick.


when i finish a stack, i grab it like this and move it to the side, then line up a new stack of 20 plates:

moving a stack of finished plates out of the way


about 30-50 plates in (depending on how thin i poured it) my first bottle of 500ml runs empty, so i repeat the alcohol wipe down and use the strap wrench to break it open before removing the lid gently by hand:

wiping down the second bottle


then i straighten up the stack again and resume the pattern: lift lid + stack, pour 1/3-1/2 plate, replace plate, lift again with the lid in place and tilt back and forth until there  are no more bare spots. i do this until the agar starts thickening up and i can see disturbances in the surface of the agar (where clumps distort what we want to be a level 2d field of media). at that point i revert back to the kosher method of pouring the entire plate with no lifting/tilting involved

pouring the rest of the plates


now i let these sit until they have set (normally the first ones are done setting and ready to be used by the time i finish the last ones), and use immediately OR bag up for future use. since ziploc bags are sterile out of the box, i like to use ziploc freezer bags to store ready-to-go plates until they are being used. this isnt normally more than a few hours to a day in my case though. these really thin plates are not suitable for storing for any long period of time and should be used asap (they will dry out)

Parafilm Preparation:


i absolutely LOVE parafilm. when i first got some in a yeast culture kit, i had no idea how it was supposed to be used, so i stretched out the 2 foot segment that came with my kit and promptly realized i had wasted it lol.... only a few years later did i realize the way parafilm is supposed to be used, and that everything i had initially had in mind was a HUGE waste of parafilm. a little bit goes a LONG way. alternatively, you can use saran wrap (bodhi prefers it), but i love the convenience of parafilm.

i like to keep a stack of prepared parafilm wraps ready to go, beside my scalpel, as this greatly speeds up my agar workflow. to set this up, i pull a few feet length of parafilm from the dispenser (the 4" wide kind), and fold it in half to determine the middle, and cut it in the middle (leaving half attached to the roll). next, use paper clips or clamps to fasten the piece you cut on top of the other, so that it is 2 layers deep and perfectly lined up. after that, i use a retractable scalpel to cut off 1/2" to 3/4" horizontal lengths, 2 at a time, and form a stack with the easy-peel ends all facing the same way. as i move up the length, i readjust the clips as needed

#1 cut a length in half, #2 clamp it 2 layers thick, #3 cut off sections 2 at a time


Working With Plates:



this is how i like my workspace to be set up, with scalpel tray, alc burner and butane bunsen,
stack of parafilm strips, 2 different recipes of plates, and a box of ziplocs. ready to work.


Preparing the Scalpel:
when i start working with the plates, i like to begin each session by opening a fresh scalpel handle from one of those foil envelopes (this is optional, flaming works fine too) and take out a fresh individually wrapped #11 scalpel blade. i like to make a small stand out of the aluminum foil from the envelope, where one end is rolled up to hold the blade end at an elevation that it is not touching anything. i put this improvised scalpel stand on the far right of my workspace

getting a scalpel handle and #11 blade ready, and making an improvised scalpel holder out of the foil envelope


EDIT: I have since upgradedto avoid open flames around so much alcohol vapor. I got a Bacticinerator and I couldnt be happier with it. Gets scalpels red hot in about 5 seconds and frees up both hands for labeling


this brings me to my flame setup. i absolutely love this Blazer brand butane bunsen burner. this japanese brand rocks in general (their torches, not their fuel), and i have for a long time been a fan of their Big Shot GT8000 for concentrates (it gets a ti nail red hot in less than 10 seconds, everyone who sees it goes out and buys one, its like a fucking lightsaber of flame haha). the butane bunsen burner is AWESOME for our purposes because it gets the scalpel red hot in about 2 seconds (it is also PERFECT for a hands-free quartz nail torch, so you can prep your dab while it is heating!!! truly awesome :rockon: ) . the only real downside is the piezo ignition, which (like all of these type igniters) will eventually wear out with too much use. i have been through 3 GT8000s and still own 2 (1 stolen, 2 ignition failed, 2 still work), and i have worn the ignition out on my first bunsen burner. for this reason, i like to use a combination approach, where i use an alcohol lamp on very low just to ignite the butane as needed. if i was only doing a few sleeves of plates at a time, i would probably just leave the flame on the whole time, but it would heat up after too long so i play it safe and just run it long enough to flame the scalpel as needed

using a butane bunsen burner and an alcohol lamp to flame a scalpel blade


next i take out a plate from the container/pile/stack to be transferred, and remove it from its individual plastic bag (more on this later), remove its parafilm wrapper, and visually inspect the culture (sometimes aided by a jewlers loupe or magnifying glass). to get a good look, i like to keep a light under my workspace rack which i turn on for backlight as needed, and i also hold the culture up to the light to get a good look

inspecting a culture in various lighting


ok so the kosher way of doing this next part (taking the sample to transfer) is to leave the plate on the worksurface, lift the lid slightly, and quickly slice out a sample 1-4mm square from the leading edge of the myc you want to transfer. i like to do something a little dicey, especially if i have cause to be concerned. i like to remove the lid, then pick up the dish and tilt it so that light reflects off of the surface of the agar. this allows us to see patterns, distortions, and blemishes which are otherwise invisible to us, and can help you spot a colony of bacteria that you might have otherwise unknowingly transferred with your myc. if you are going to do this, its important that you have slick, even, smooth agar to begin with, otherwise you cant really draw any conclusions from distortions in the reflective surface. unfortunately i couldnt get a good shot of what this technique of spotting hidden contams looks like, but hopefully yall get the picture

using reflecting light to identify hidden contams and irregularities in the agar surface while taking a sample


after that, we make the transfer to a fresh plate and label it with a sharpie (black for MS, red or blue for clones or isolates). i usually work with stacks of 8 plates at a time and start with the top and work down, opposite of pouring.  next we lift the lid slightly and then place the tissue sample onto the center. in the past, i used to stab the agar right through the center of my transfer tissue, as if to "nail it in place", but quit doing this for several reasons. mainly, it causes the growth pattern to be elongated around the blade trench (rather than uniformly circular), and also it doesnt do a good job of holding the sample in place (often makes it worse). it might seem a little counter-intuitive, but all you need to do is lay the piece of tissue down on the receiving dish, whether it is right side up, upside down, on its side, or whatever, does not matter. the surface tension between the two agar surfaces (or the myc and the agar if its upside down) will be more than sufficient to hold it in place, and the growth will be regular. if you try to readjust it a bunch of times or stab it or something, it can slide or fly all over the place, or even split into pieces, and make it hard to draw any conclusions from the shape of the resulting colony. Just dont drop your plates!

transferring a sample to the receiving dish


now i wrap the receiving dish in parafilm. first, i grab a piece from the pile and peel off the paper backing. next i pinch one end between my index finger and the dish, and rotate the dish thereby stretching the parafilm around the outside edges of the dish.

sealing a dish with parafilm


i like to put each plate into an individual ziploc sandwich bag, since they come sterile and it makes them more durable and easier to handle and inspect outside a sterile environment

individual ziplocs for each plate


Working with Spores:

since we fruit in open air, we can assume that spores are dirty. if someone sends you a spore print, you can assume it is going to have contams on it in addition to the spores you want to cultivate. this does not mean you got a bad print, it means you got a NORMAL print. the same thing applies to syringes. for this reason, when inoculating something with spores, it is a good idea to use as little as possible, since we only need a few spores and the more solution/spores we use, the more contams will be present as well. i recommend people never putting spores directly to grain, since it is so easy to culture the contams

a sterile jar of grain is basically the microbiological jackpot for anything that can digest it (fungi, bacteria, etc), so we need to take care to reduce the chances of culturing the wrong microorganisms. basically, ALWAYS germinate your spores first on agar before applying them to grain/lc/etc.

by using streaking techniques, you can easily clean up even the dirtiest of prints or spore syringes. you basically are using a loop to spread spores VERY thin across the surface of agar, creating various zones with different concentrations of spores. this is extremely useful for cleaning up a print, since you spread the contams thin while you spread the spores. this is also a great way to obtain isolates about 100 transfers sooner than you would get from a blob of spores, since the later zones have very few strains present in each colony. by doing one of these serial dilution style streaks, you end up with hundreds of colonies to choose from, of various genetic diversity, which gives you lots of chances to select a vigorous, organized culture

Quote:

In microbiology, streaking is a technique used to isolate a pure strain from a single species of microorganism, often bacteria(fungi in our case). Samples can then be taken from the resulting colonies and a microbiological culture can be grown on a new plate so that the organism can be identified, studied, or tested.

Streaking is rapid and ideally a simple process of isolation dilution. The technique is done by diluting a comparatively large concentration of bacteria to a smaller concentration. The decrease of bacteria should show that colonies are sufficiently spread apart to effect the separation of the different types of microbes. Streaking is done using a sterile tool, such as a cotton swab or commonly an inoculation loop. Aseptic techniques are used to maintain microbiological cultures and to prevent contamination of the growth medium.There are many different types of methods used to streak a plate. Picking a technique is a matter of individual preference and can also depend on how large the number of microbes the sample contains.

The three-phase streaking pattern, known as the T-Streak, is recommended for beginners.The streaking is done using a sterile tool, such as a cotton swab or commonly an inoculation loop. The inoculation loop is first sterilized by passing it through a flame. When the loop is cool, it is dipped into an inoculum such as a broth or patient specimen containing many species of bacteria. The inoculation loop is then dragged across the surface of the agar back and forth in a zigzag motion until approximately 30% of the plate has been covered. The loop then is re-sterilized and the plate is turned 90 degrees. Starting in the previously streaked section, the loop is dragged through it two to three times continuing the zigzag pattern. The procedure is then repeated once more being cautious to not touch the previously streaked sectors. Each time the loop gathers fewer and fewer bacteria until it gathers just single bacterial cells that can grow into a colony.The plate should show the heaviest growth in the first section. The second section will have less growth and a few isolated colonies, while the final section will have the least amount of growth and many isolated colonies.




examples of streaked plates. different labs use different standards for streaking. colonies thin as the dish rotates counter clockwise


the way i like to do it is very similar to what is described above. basically, take your loop (bod's DIY loop tek) and scrape off a small amount of spores from a print (or drip a drop from a syringe directly onto the loop) and streak a side of the plate like below. then flame the loop again, DONT SCRAPE MORE SPORES, rotate the dish, and drag the loop from the area you streaked previously to streak zone 2, then flame and repeat

this is the streaking technique i like to use. it is tremendously handy for cleaning up spore prints, isolation projects, and even cleaning up old LCs


if you are using a spore syringe as your starting point (or a LC you are trying to refresh), you want to be super careful not to add any additional liquid to the plate, otherwise it will roll around and distort the streaking pattern. so make sure you dont drip over the plate, you shouldnt be putting any more liquid into the plate than what sticks to the loop

Conclusion:


so that is pretty much it!! a bit lengthy, but i think i covered most of what i wanted to cover for my post #1000. please let me know if yall have any questions or need clarification about anything :smile:

i also welcome any additional tips & tricks, feedback, criticism, discussion, debate, haters (:kiss:), or critical thinking :wink:

thanks everyone for everything yall have done to make this place such an incredible resource. im so grateful to live in an age where there is an online community to collaborate with on our hobbies :smile:

warm regards,
c10


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C10's Agar Guide + Tips and Tricks | c10's Flow Hood Build Guide


"Partial knowledge is more triumphant than complete knowledge; it takes things to be simpler than they are, and so makes its theory more popular and convincing."

"Convictions are more dangerous enemies of truth than lies"
― Friedrich Nietzsche


Edited by c10h12n2o (02/14/20 05:27 PM)


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: c10h12n2o] * 3
    #24330963 - 05/18/17 09:01 PM (3 years, 4 months ago)

:tldr:

but i will. this is awesome. happy 1000


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: mushboy] * 1
    #24330989 - 05/18/17 09:08 PM (3 years, 4 months ago)

:superscience:      :hi:      :takingnotes:


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: mushboy] * 2
    #24330997 - 05/18/17 09:09 PM (3 years, 4 months ago)

Congratulations on post 1000! Great write up and  well documented with pics.  That's a +5  :mushroom: for ya! Keep the good stuff coming.


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: Just_A_Noob] * 1
    #24331001 - 05/18/17 09:11 PM (3 years, 4 months ago)

So this is why you've been so quiet lately!
What a post dawg!

Very good sir.
I concur.
Grate  write up, lots of info in once place

(keywords:agar, agar magic, amazing agar, easy agar, flow hoods, flowhoods, ahgar, Petri, plate, amazeballs, parafilm, recipe, suspension, incredible)


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: mynakedrat] * 1
    #24331014 - 05/18/17 09:18 PM (3 years, 4 months ago)

lol we should put you on SEO duty MNR :lol:

thanks guys :smile:

yeah this is why ive been so quiet. i got to post 99X and didnt have the guide ready yet so i was like damn... cant start any shit until i get this posted
:cryaboutit:

also my camera messed up right before i made this, so i had to use my backup camera instead of the one i bought for photographing this kind of stuff :/ its being repaired right now and should be back before too long


--------------------

C10's Agar Guide + Tips and Tricks | c10's Flow Hood Build Guide


"Partial knowledge is more triumphant than complete knowledge; it takes things to be simpler than they are, and so makes its theory more popular and convincing."

"Convictions are more dangerous enemies of truth than lies"
― Friedrich Nietzsche


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: c10h12n2o] * 1
    #24331233 - 05/18/17 10:52 PM (3 years, 3 months ago)

As usual, great info my friend. Well done


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OfflineKenetic
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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: Pipefitter537] * 1
    #24331269 - 05/18/17 11:07 PM (3 years, 3 months ago)

Wow! What a comprehensive and useful tek! I can't read it all tonight but be assured I will read it all. 

You are obviously very skilled.

Happy 1000th!!!  :birthday:  :birthday:  :birthday:  :birthday:  :birthday:  :birthday:  :birthday:  :birthday:  :birthday:  :birthday:  x  100


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: Kenetic] * 1
    #24331380 - 05/18/17 11:50 PM (3 years, 3 months ago)

Great stuff sir!  Nicely illustrated too.

A big take-away for me is to buy a strap wrench, what a clever invention... no more battles with stubborn mason jar lids!

I will buy one tomorrow.  YASSSSSSS :thumbup: :smile:


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: mushboy] * 1
    #24331393 - 05/18/17 11:56 PM (3 years, 3 months ago)

Quote:

mushboy said:
:tldr:

but i will. this is awesome. happy 1000




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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: Lobi] * 1
    #24333334 - 05/19/17 05:30 PM (3 years, 3 months ago)

much obliged my friends

:fuckyeahdance:

ive been planning on doing an agar tips thread for a long time, but figured since i do so many plates it might be a good idea to spell out my whole process and show what i have learned as far as what works and saves time on that scale (did 100-200 plates a day for several months lol). plus it gives people the chance to correct me on any dumb shit i do/think/believe :lol:

there are a few things i will add better pics of when i get my camera back from the shop (used my phone for all these :/ ), specifically the technique using light reflections to spot hidden contams on agar

i will try to update this to make it reflective of what i am currently doing as time goes on and i revise my technique :smile:


--------------------

C10's Agar Guide + Tips and Tricks | c10's Flow Hood Build Guide


"Partial knowledge is more triumphant than complete knowledge; it takes things to be simpler than they are, and so makes its theory more popular and convincing."

"Convictions are more dangerous enemies of truth than lies"
― Friedrich Nietzsche


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: c10h12n2o] * 1
    #24334504 - 05/20/17 01:29 AM (3 years, 3 months ago)

I love your crystal-clear agar plates! :congrats:


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: c10h12n2o] * 3
    #24334565 - 05/20/17 02:02 AM (3 years, 3 months ago)

Just reading this made me feel smarter.


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: WeavieWonder] * 1
    #24334572 - 05/20/17 02:05 AM (3 years, 3 months ago)

Great stuff. I has questions.


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: Kingdeviluke] * 1
    #24334657 - 05/20/17 02:57 AM (3 years, 3 months ago)

Tonight was the first time i used a print ive only used syringes... say the prints just show bacteria how would you go getting clean growth ?


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: Boogieman47] * 2
    #24334758 - 05/20/17 04:25 AM (3 years, 3 months ago)

fire away, all questions/comments welcome :smile: I appreciate yells feedback!

Quote:

Boogieman47 said:
Tonight was the first time i used a print ive only used syringes... say the prints just show bacteria how would you go getting clean growth ?




nice! I bet you'll get hooked and end up with a box or photo album full of prints (a lifetime supply of genetics to work with, in as many flavors as you like) to pull from as needed


I'm actually glad you asked that. I'm in the shower right now but I will update the guide to include my streaking techniques

but to answer your question: I would do a serial dilution streak, and watch it closely. in 99/100 cases, that will be more than sufficient to separate from contaminants, just by streaking and making well-timed transfers. alternatively you could just do a single dilution and streak, by scraping a tiny bit of spores onto a loop then dilute them into a tiny bit of sterile water, then just do a single streak pattern. either works great, but I prefer the method without water.


if that didn't work (it almost always does) I'd streak antibiotic agar


--------------------

C10's Agar Guide + Tips and Tricks | c10's Flow Hood Build Guide


"Partial knowledge is more triumphant than complete knowledge; it takes things to be simpler than they are, and so makes its theory more popular and convincing."

"Convictions are more dangerous enemies of truth than lies"
― Friedrich Nietzsche


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: c10h12n2o] * 1
    #24334775 - 05/20/17 04:42 AM (3 years, 3 months ago)

I just got some wire made a small loop and flamed it cooled in the agar and streaked the whole in a zigzag somewhat like that gif....

I have about 10 prints pans and cubes i have made tons of prints to send out but figured in my arrogant mind i wouldnt need to work with them lmao i ordered 14 syringes over a year ago 6 unopened and the rest are pretty full maybe 2 cc's if that ...


On each plate i dropped 5 drops in a 5 point star 2 plates per syringe 4 kinds and made 3 plates of prints with 3 varieties ...


I guess i was thinking a print will just be a booger of bacteria like the syringes sometimes do haha ..


Have you worked with black tea agar or charcoal?? Also my buddy has a silver water machine and said its antibacterial i was wondering if i was to make spore solution with it if it would help with cleaner syringes


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: Boogieman47] * 1
    #24334839 - 05/20/17 05:48 AM (3 years, 3 months ago)

When tranfering how do you not make the agar stick to the hot knife if you don't stab the plate


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: Kingdeviluke] * 1
    #24336286 - 05/20/17 07:43 PM (3 years, 3 months ago)

Quote:

Boogieman47 said:
I just got some wire made a small loop and flamed it cooled in the agar and streaked the whole in a zigzag somewhat like that gif....



loops are awesome, totally indispensable. for a long time i used to streak agar, but didnt really understand what i was doing. i would just do a basic streak pattern. at some point i made the connection that streaking is actually smearing something really thin, and then learned how to do the serial dilution streak like the gif. the biggest difference is the serial dilution streak involves scraping spores 1 time, streaking a side, then flaming, then smearing from the area you streaked before into a new area, then flaming again and repeating. the key is that you only scrape spores for zone 1, and each progressive zone has less and less spores per area. this is a really easy way to get isolates and highly organized cultures much faster (since anything from zone 4-5 is gonna have way less genetic diversity). you end up with something like this (sorry i dont have a more clear pic handy)


Quote:

Boogieman47 said:
I have about 10 prints pans and cubes i have made tons of prints to send out but figured in my arrogant mind i wouldnt need to work with them lmao i ordered 14 syringes over a year ago 6 unopened and the rest are pretty full maybe 2 cc's if that ...



i know the feeling man. thats why i made the album of my prints, so that i keep at least 1 of each and have the origin documented, etc. i keep extras in a box, in ziplocs, and send those out for trades. greatly reduces the risk of accidentally sending out my last one

when you streak like this, it really doesnt take hardly ANY spores... when i do a trade im happy if someone just gives me a swab of part of a print lol... its so easy to turn a few spores into as many prints as you want :smile: prints seem like the best long term way to store things. i dont trust syringes at all, even though ive had them last many years, there are just too many things that can go wrong, where with a print you can do whatever you want with it. really, a print is a lifetime supply for me. they are microscopic, we waste so many haha...

i would try to grow out all your syringe varieties (streak to agar > a2g or LI > grainspawn > mono) and make prints for long term storage. while you are at it, make slants out of any clones you particularly like :smile:

Quote:

Boogieman47 said:
On each plate i dropped 5 drops in a 5 point star 2 plates per syringe 4 kinds and made 3 plates of prints with 3 varieties ...



that sounds risky! when i am working with a syringe, i like to drip the solution directly onto my loop, then streak with that in the gif pattern (flaming between each step). you could also drip some into a sterile shot glass or something and run it through that. the point is that you only need a microscopic amount of spores, and the less the better.

since we can basically assume spores are dirty (whether print or syringe), the less we use the better. and if we can smear that tiny amount across the agar to thin it out even more, even better. also, we dont want to add ANY liquid to the plate if we can help it, because this will roll around and inoculate random spots with spores and contams, making it so that we cant infer anything from the shape. if we just use as little as possible, then it makes it easier to separate the good colonies from the bad since they remain distinct. any liquid on the agar surface is going to cause problems, so keep it to a minimum and spread out the contents

Quote:

Boogieman47 said:
I guess i was thinking a print will just be a booger of bacteria like the syringes sometimes do haha ..



thats a pretty good assumption to be honest :lol: they damn sure can. i assume spores=dirty. some prints are insanely dirty, just like some syringes (which are made from prints). id prefer a dirty print over a dirty syringe any day though, because in a syringe the liquid spreads the contams everywhere and gives them the h2o they need to reproduce. its much easier to work with a given amount of spores from a print, wheras i have no idea how many they put in a syringe. same process applies to cleaning up either one though: streaking and well-timed transfers

Quote:

Boogieman47 said:
Have you worked with black tea agar or charcoal?? Also my buddy has a silver water machine and said its antibacterial i was wondering if i was to make spore solution with it if it would help with cleaner syringes



i havent. i did experiment with my agar recipe a lot, but i eventually realized that i was asking the wrong questions (thinking about antibiotics, nutrition, etc). with agar all you need is a slightly nutritious medium that allows for growth in 2 dimensions at a controlled rate. trying to "optimize" the nutrition completely misses that point.

i do use antibiotics occasionally, but very very rarely. 99/100 times, well timed transfers and basic MEA or PDA is all you need to clean up a print, and if you use a streaking technique it can be done in 1 transfer from pretty much anything, no matter how dirty. the only time i use antibiotics is occasionally on the final plate before making long-term master slants (the final plate, not the slant itself, for that i use standard full strength MEA), and if a print is so filthy that the bacteria is keeping the spores from germinating

id say the better question to ask would be why you want to be working with syringes at all? all i use them for is sucking up LI. i doubt i will ever touch a spore syringe again, besides what i get in trades, etc.. what exactly do you have in mind for syringes that couldnt be more effectively done with agar and g2g? i could very well be missing something :lol:

Quote:

Kingdeviluke said:
When transferring how do you not make the agar stick to the hot knife if you don't stab the plate




good question, i need to make that more clear in the guide. we are talking about 2 different things: in the guide i was referring to after i take the transfer and set it into the receiving dish. i used to (and sometimes still do from habit) place the transfer on the receiving dish and then stab the transferred tissue, piercing it and the agar in the receiving dish, as if to nail it in place. this often caused the transferred tissue to split, and distorted the colony growth pattern because of myc cells on the blade, which often cause the colony to grow out of the gash made by the blade. these days, instead of that, i try to just sit it down and move on, since the surface tension holds it just fine and the stabbing doesnt really help

what you are asking about is something different. i heat the blade red hot in the butane bunsen burner and then set it down onto that little aluminum foil tray i make each session, which holds it at an angle so that the blade doesnt contact anything. i time things so that as soon as i make a transfer the next thing i do is flame the scalpel, then put it down on that tray, then i label, parafilm, and bag the donor and recipient plates, and get out the next donor plate and unwrap it. by the time i am done with all that, the blade is always cool enough to do the next transfer without problems.

the exception to this is when i have a plate that i am taking multiple transfers from, since that doesnt give the blade long enough to cool down between transfers. in that case, i do drag the hot blade through the receiving dish to cool it down. this is also a handy way for me to id cultures, since i know that any plates that have that scar from cooling the scalpel were done in a series from the same donor

does that make sense guys? let me know if yall have any questions about anything :smile:


--------------------

C10's Agar Guide + Tips and Tricks | c10's Flow Hood Build Guide


"Partial knowledge is more triumphant than complete knowledge; it takes things to be simpler than they are, and so makes its theory more popular and convincing."

"Convictions are more dangerous enemies of truth than lies"
― Friedrich Nietzsche


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Re: c10's Agar Guide + Tips & Tricks (post #1000) [Re: c10h12n2o] * 1
    #24336637 - 05/20/17 10:20 PM (3 years, 3 months ago)

I like the distinction you make into the differences of using a flowhood in comparison, and contrast to a SAB.
The different color markers representing the different types of culture is cool, too.
You're a hoss for picking up all 19 plates at a time to pour.
I need to get me a nice swivel chair, and some lighting like your's for my DIY lab/clean room, and spawning area.

Excellent write up!  :super:


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