Helminthic Therapy wiki - User contributions [en]

User talk:Alana

2018-04-20T19:31:37Z

Alana: /* Skin Infections */ new section

==Alana meet Tracy==

Hi Alana, another wiki User (Tracy) wants to put a couple of questions to you about your NA and TTO protocols, so I’ve suggested that she uses the wiki to do this because it allows wiki Users to communicate with one another via the email addresses they signed up with, without actually revealing those addresses to anyone. Only the wiki software knows what these are.

I’ve sent a message to Tracy, telling her what to do to contact you this way, and you should be able to read my message to her here: http://helminthictherapywiki.org/wiki/index.php/User_talk:Tracy You can also write on her Talk page, if you want to, after hitting the “Edit” button at the top right of the page.

Once you've finished your message, add four tildes (~), which the wiki will replace with your name, as a signature, after you’ve hit the “Save page” button.

If Tracy sends you a message using your Talk page, you can reply to her on this page, after hitting the "Edit" button.

If you want to remove any messages, once they've been seen, just re-enter the page using the "Edit" button, highlight/delete the comment, and hit "Save page". [[User:John|John]] ([[User talk:John|talk]]) 03:48, 18 September 2017 (UTC)

== Clarification of Your Incubation method for TTO and NA ==

Hello.

I have read through your TTO protocol (thank you very much for creating such detailed document for the wiki) and just need a bit of clarification about your incubation protocol.

Question 1 - I understand that many use Sarah's method for incubating NA but would like to know what your practice is, please.

Question 2 -have you ever found a research paper about incubating TTO? I suspect not as you have included it in your thorough cited sources. I just wanted to double check.

I am going to attempt to incubate and process my own TTO and was hoping I could get this bit of clarification.

My background. I have been using TSO and NA and TTO to keep Crohn's Disease in remission for the past 11 years and have experimented with them to see what works best and TTO and TSO work best for me. TSO is prohibitive because of its cost and fortunately, TTO has worked very well for me. I also use NA for its systemic effect and I think it works well as an added buffer against autoimmune response activation, in general.

John Scott suggested I contact you. So here I am. Thank you in advance for your suggestions.

[[User:Tracy|Tracy]] ([[User talk:Tracy|talk]]) 19:17, 2 October 2017 (UTC)

==Testing==

I'm checking to see whether you get a notification of this message, Alana.

[[User:John|John]] ([[User talk:John|talk]]) 02:30, 3 October 2017 (UTC)

Hi John. I'm responding to your message. I received an email notification. [[User:Tracy|Tracy]] ([[User talk:Tracy|talk]]) 16:33, 3 October 2017 (UTC)

== Hello John and Tracy ==

This is just confirming that I'm now able to use this forum.
[[User:Alana|Alana]] ([[User talk:Alana|talk]]) 21:20, 13 October 2017 (UTC)

Hi Alana & John! This is wonderful.

Alana, Thank you for your input so far. Your protocol is wonderfully detailed. I am currently making a list of all the materials I will need and referencing it with the materials at the lab.
Fortunately, the bio-hackers that formed and created the lab space are keen adventurous university grad students and researchers at a local hospital, making access to reagents and other medical grade supplies possible.

It should take couple more weeks to get underway, I need to order a few things - 20 micron nylon mesh is expensive!

Hydrometer question: Can I use a hydrometer for brewing? I would like to if I can because I brew kombucha. https://www.amazon.com/dp/B0735B5YND?psc=1

[[User:Tracy|Tracy]] ([[User talk:Tracy|talk]]) 21:49, 13 October 2017 (UTC)

== Chefast Hydrometer ==

Since I haven't used this brand of hydrometer before I needed to take a quick look at the protocol and be sure that this hydrometer covered the full range of densities discussed in the paper. If it did then I don't see that there should be any problems. So https://www.amazon.com/dp/B0735B5YND?psc=1 says, "Measuring specific gravity from 0.99-1.17" Sorry, but you need to be able to go up to 1.270 for the Sheather's solution. In the paper I list "Wards Natural Science Specific Gravity Hydrometer 1.00 - 2.00 model." (I also just now realized I mistakenly included one that wouldn't have worked. "150879 Specific Gravity Hydrometer, 1.00–1.20, 0.002 divisions (see graphic 2J)") [[User:Alana|Alana]] ([[User talk:Alana|talk]]) 22:08, 13 October 2017 (UTC)

I'm glad I checked. The "Wards Natural Science SGH 1.0 - 2.00 it is!
Thank you. [[User:Tracy|Tracy]] ([[User talk:Tracy|talk]]) 22:14, 13 October 2017 (UTC)

==Hydrometer selection==

Alana, would you like to remove the unsuitable hydrometer from the materials list in your protocol? If you click on the "[Edit]" button next to the heading, "Lab Equipment", you'll be able to locate the relevant text in the source code and remove it. [[User:John|John]] ([[User talk:John|talk]]) 23:07, 13 October 2017 (UTC)

==Editing MediaWiki markup==

Here are some references to help you with editing the site. The Cheatsheet covers most of the things you're likely to need, and you may be able to find examples of things you want on other pages on the site. If you hit a problem with the markup, just message me on my Talk page: http://helminthictherapywiki.org/wiki/index.php/User_talk:John

Cheatsheet: https://en.wikipedia.org/wiki/Help:Cheatsheet 
The simplest possible guide to writing MediaWiki: https://www.sjbaker.org/wiki/index.php?title=The_simplest_possible_guide_to_writing_MediaWiki 
The Missing Manual: Editing for the first time: https://en.wikipedia.org/wiki/Help:Wikipedia:_The_Missing_Manual/Editing,_creating,_and_maintaining_articles/Editing_for_the_first_time 
Help:Formatting: https://www.mediawiki.org/wiki/Help:Formatting 
Links: https://www.mediawiki.org/wiki/Help:Links 
Lists: https://www.mediawiki.org/wiki/Help:Lists 
Images: https://www.mediawiki.org/wiki/Help:Images 
Citations: https://www.mediawiki.org/wiki/Extension:Cite 
Quote box: http://stackoverflow.com/questions/131049/whats-the-easiest-way-to-add-a-quote-box-to-mediawiki 
Tables: https://www.mediawiki.org/wiki/Help:Tables 

[[User:John|John]] ([[User talk:John|talk]]) 22:56, 13 October 2017 (UTC)

== Evil Hydrometer Removed! ==

Thanks John. That process went quite well. [[User:Alana|Alana]] ([[User talk:Alana|talk]]) 18:35, 14 October 2017 (UTC)

== Clarification on larvae sterilization ==

Hello, I was just reading about incubating and had a few questions.

"The larvae are forced to migrate through a 3 inch layer of activated charcoal"? 3 inches is a lot! Is that really activated charcoal and not just normal charcoal?

"they are removed by placing two layers of fabric"? like an old t-shirt? the actual incubating instructions mentioned gauze, but I guess fabric is better since it can be washed and reused and doesn't have to be cut and carefully laid each time

"The sample is washed in buffered saline solution by agitating them mechanically for twenty minutes. The larvae are extracted from the saline by pipetting them from the surface (they float in saline) and are then placed on the surface of a piece of filter paper. They are submerged and flushed ten times using a solution of Chlorohexidine 0.2% by weight."
So are they washed in saline solution, Chlorohexidine, or the buffer solution linked below, because the buffer solution looks like a huge effort to prepare! 3 inches of charcoal and flushing 10 times also sounds a bit over the top. And how can you even be sure that you pipette all of them up when you can't even see them??

Is any of that really necessary? What's the bare minimum you absolutely need to do to insure safety when giving NA to someone else, as far as supplies, process, and blood/stool tests?

[[User:Mäuschen|Mäuschen]] ([[User talk:Mäuschen|talk]]) 21:25, 18 April 2018 (UTC)

== Skin Infections ==

While a skin infection always is a possibility, I've been self-inoculating for about 10 years and haven't had any infections. For the first 5 years I was inoculating every 6 months, and for the last 5 I've been inoculating every year, which adds up to about 15 inoculations. Using the Alana protocol, the NA crawl up through the soil, gauze, and into the cork. Then I've simply been collecting the larvae from the cork, storing them in water, and then using them after sterilizing my skin with alcohol (and then letting it dry). I've never used any of the materials or methods you mention.

These organisms have been infecting people for probably millions of years, and they have evolved to have a symbiotic or at least a commensal relationship with their host. The reasoning is if they harmed their hosts then their genes would be selected against. So they have evolved, at least somewhat, to avoid doing this. For example, they shed their outer layer of skin when they enter the upper layer of your skin, which also sheds the bacteria that might be on them.

Although there could be reports of infections I'm unaware of, I've never heard of this happening to anyone in the community. You might check with others in the DIY culturing community and see what their experiences have been. And I've never run across any reports in journal articles or texts about people who live in underdeveloped areas having such issues. But perhaps I simply haven't run across them, or such infections haven't been reported from the field. So I don't know what the actual odds of an infection are. If you can't find any reports in the helminthic community or the related literature, I suppose the next thing to look at for estimating the risks would be looking at various closely analogous species of helminths that frequently infect non-human animals, and see what the relevant veterinarian literature might say about the possibility of skin infections.

I suspect that these companies have to be extremely careful because they're justifiably concerned about liability. If something ever did go wrong, even coincidentally, think about what it would be like to go into court and try to defend yourself. You'd want all the ammunition you could get.

[[User:Alana|Alana]] ([[User talk:Alana|talk]]) 19:31, 20 April 2018 (UTC)

User talk:Tracy

2017-10-15T19:30:34Z

Alana: /* Hookworm Culturing before TTO */ new section

==Chatting in the wiki==
Tracy, if you want to talk to another wiki User (editor), like I'm talking to you here now, go to the list of users (http://helminthictherapywiki.org/wiki/index.php/Special:ListUsers) and click on "talk" beside that User's name. This will open their "talk" page. Write your message in the blank space on the page and then add 4 tildes (~) which the software will replace with your user name once you hit the "Save page" button at the bottom. The message will then be sent to whatever email address that User signed up with.

This communication method will allow you to talk directly to Alana, for example, about her two incubation protocols in the wiki, or to any other User about any other wiki matter! How cool is that?

To make a change to any wiki page, just click on the "Edit" button at the top right of the page, and hit "Save page" when you're done - or "Show preview" if you want to check any details before saving. You'll soon get the hang of it! [[User:John|John]] ([[User talk:John|talk]]) 03:18, 18 September 2017 (UTC)

Hi John, My first attempt at communicating through "talk."

It might be worth bookmarking the list of Users: http://helminthictherapywiki.org/wiki/index.php/Special:ListUsers This will give you access to all the others.

And now you can ask Alana about her incubation protocols. Here's her Talk page: http://helminthictherapywiki.org/wiki/index.php/User_talk:Alana As you'll see, I've alerted her to the fact that you'll be getting in touch! [[User:John|John]] ([[User talk:John|talk]]) 14:57, 18 September 2017 (UTC)

Thank you, John! I left a message with Alana. Fingers crossed.

==Editing MediaWiki markup==

Here are some references to help you with editing the wiki. The Cheatsheet covers most of the things you're likely to need, and you may be able to find examples of things you want on other pages on the site. If you hit a problem with the markup, just message me on my Talk page: http://helminthictherapywiki.org/wiki/index.php/User_talk:John

Cheatsheet: https://en.wikipedia.org/wiki/Help:Cheatsheet 
The simplest possible guide to writing MediaWiki: https://www.sjbaker.org/wiki/index.php?title=The_simplest_possible_guide_to_writing_MediaWiki 
The Missing Manual: Editing for the first time: https://en.wikipedia.org/wiki/Help:Wikipedia:_The_Missing_Manual/Editing,_creating,_and_maintaining_articles/Editing_for_the_first_time 
Help:Formatting: https://www.mediawiki.org/wiki/Help:Formatting 
Links: https://www.mediawiki.org/wiki/Help:Links 
Lists: https://www.mediawiki.org/wiki/Help:Lists 
Images: https://www.mediawiki.org/wiki/Help:Images 
Citations: https://www.mediawiki.org/wiki/Extension:Cite 
Quote box: http://stackoverflow.com/questions/131049/whats-the-easiest-way-to-add-a-quote-box-to-mediawiki 
Tables: https://www.mediawiki.org/wiki/Help:Tables 

[[User:John|John]] ([[User talk:John|talk]]) 22:59, 13 October 2017 (UTC)

== Hookworm Culturing before TTO ==

Hi Tracy,

Earlier you asked about the advisability of skipping the hookworm culturing step, and I didn't see any likely problems with that. After thinking about this I thought I should add a few comments.

The NA eggs should hatch within about 24 hours into stage 1 larvae. These aren't infectious, and it should take 5 days before there could be any stage 3 infectious larvae in the stool. The worst case scenario I can imagine would be if the larvae in the Sheather's solution could somehow survive eating stool in that mixture, while floating to the top of it, so you transferred them along with the eggs you were cleaning. Then by day 5 they had molted into infectious stage 3 larvae. Then when you are handling the materials you accidentally get a large infection. When I write this down it all sounds very implausible, but whenever you change something there's always at least the theoretical possibility something unexpected could happen. I don't know if stage 1,2,or 3 larvae will float in Sheather's, and I don't imagine they can survive under those conditions, but I've never tested any of this. [[User:Alana|Alana]] ([[User talk:Alana|talk]]) 19:30, 15 October 2017 (UTC)

User talk:Alana

2017-10-14T18:35:04Z

Alana: /* Evil Hydrometer Removed! */

User talk:Alana

2017-10-14T18:34:43Z

Alana: /* Evil Hydrometer Removed! */ new section

TTO incubation: very detailed method by Alana

2017-10-14T18:31:28Z

Alana: /* Lab Equipment */

{{Breadcrumb|[[Helminth_incubation|Incubation]]}}
<div class="toclimit-3">__TOC__</div>
== Introduction ==
Having experienced considerable benefit to my health as a result of hosting a controlled number of human
whipworms ([http://en.wikipedia.org/wiki/Trichuris_trichiura Trichuris trichiura]), I wanted to be able to maintain my whipworm colony with as little expense as
possible. After carrying out a search of the literature, followed by experiments to refine the process, I arrived at the
protocol set out below.
The next section, “Warning and Disclaimer”, is essential reading because there are significant safety, and potentially
legal, implications to helminth incubation.

== Warning and Disclaimer ==
* There is considerable risk involved in incubating whipworm eggs, due to their infectious nature. After successfully culturing whipworm eggs, it would be easy for someone to accidentally ingest many thousands of these organisms, with potentially very serious consequences for the individual’s health - for example, resulting in [http://www.manchester.ac.uk/discover/news/article/?id=5841 rectal prolapse] - and perhaps also the future of helminthic therapy if news of such an accident were to reach the media.

* It may be illegal to incubate whipworm eggs in some jurisdictions, including the U.S., where they are currently classified by the Food and Drug Administration (FDA) as biological agents (i.e. drugs), as defined in Section 351 of the Public Health Service Act, and subject to an [http://www.accessdata.fda.gov/cms_ia/importalert_159.html Import Alert].

* The information contained in this document is not advice, but for general information only. It has not been approved or evaluated by any governmental organisation concerned with the regulation of healthcare or drugs anywhere in the world. The accuracy, validity, effectiveness, completeness or usefulness of this information cannot be guaranteed.

* While the information presented here is related to the practice of the experimental treatment known as helminthic therapy, it is not intended to provide medical advice, diagnosis or treatment. The reader is hereby advised to always consult with a physician or other professional health-care provider regarding any health care problem or issue they might have.

* Anyone who chooses to make use of the information in this document does so at their own risk and no responsibility or liability whatsoever is accepted by the author for the use or misuse by others of any of the information contained in this document.

== Purpose of incubation ==

My intention is to clean and incubate the eggs produced by the colony of whipworms (Trichuris trichiura) that I am
already hosting. This will allow me to re-infect myself for the purpose of maintaining the beneficial effects I have
from harbouring them.

My existing whipworms were purchased as eggs from a recognised provider of helminthic therapy, and I'd never use
eggs from any other source because I want to be certain that any organisms I introduce into myself are either
harvested from my own body or from a donor whom I can be confident is free from pathogenic organisms and, in
the latter case, that the organisms supplied have been scrupulously cleaned.
<ref name="cleanup" group="note" />

== The laboratory and safe practice ==

* I have created a designated work area where no food or drink is prepared or allowed.
* I always wear long sleeves, long pants, socks and shoes plus a lab apron while following this protocol.
* I always wear gloves of latex or a synthetic alternative, and avoid touching my skin (e.g., face) while working.
* I use a sharps container for broken sharp objects - usually an old pill bottle. ([http://blink.ucsd.edu/safety/research-lab/hazardous-waste/sharps/ How to Dispose of Sharps])
* I use a dedicated small, lined trash container with a lid, and empty this frequently.
* I disinfect all materials using two containers. First, I dip the equipment into either undiluted 5-6% bleach or 2-3% ammonia.<ref name="Bleach" group="note" /> Then, after about 5 minutes, I clean them in soapy water, before finally rinsing and drying them. ([http://www.cdc.gov/hicpac/Disinfection_Sterilization/17_00Recommendations.html Guideline for Disinfection and Sterilization in Healthcare Facilities, 2008])
* I boil for at least 10 minutes the leftover soil and stool in my stock pot.
* I make absolutely sure my centrifuge is balanced.
* I only add acid to water, and never add water to acid.
* I neutralize my acid outside because of the caustic fumes this generates.
* I only add a little baking soda at a time when neutralizing the acid, since this generates a lot of foam.
* I disinfect all work surfaces when finished and, to be as confident as possible that nothing remains alive, I use either undiluted 5-6% bleach or undiluted 2-3% ammonia.<ref name="Bleach" group="note" /> I only ever buy and keep available one of these chemicals at a time and I store whichever one I’m using in a 0.5 litre (17 U.S. fl oz) container and apply this to the relevant surfaces, as required, using a sponge. I then wait at least 2 minutes in order to be as confident as I can be that the eggs are dead. Once the clean-up is done, I thoroughly rinse the sponges about 10 times afterwards, to prevent the residue bleach or ammonia from rapidly destroying them.
* I absolutely never mix or work with bleach and ammonia together, as to do so would produce explosive and toxic gasses.
* Whenever I use bleach or ammonia, I ensure that there is good cross-ventilation to prevent the build-up of fumes.

== Separating hookworm and whipworm eggs ==

This protocol takes into account the fact that I carry both human hookworms ([https://en.wikipedia.org/wiki/Necator_americanus Necator americanus]) and human whipworms ([https://en.wikipedia.org/wiki/Trichuris_trichiura Trichuris trichiura]). Even if I’m not planning on extracting hookworm larvae from a given culture I still culture it for 8 days, as I would with hookworms. <ref name="Robust" group="note" />
Since viable hookworm eggs hatch within one or two days, (see [[NA incubation: very detailed method by Alana | NA incubation: very detailed method by Alana]]) any such eggs will have long since hatched and their larvae will have moved on. So I’m very sure that the hookworm and whipworm eggs have been separated. (Later I’ll also be filtering the material with a 30 micron mesh, which would also remove any hookworm eggs.<ref name="TTeggs" group="note" /> I'm also confident that this understanding is correct because I've never seen any hookworms or hookworm eggs under my microscope when I've done my whipworm counting and dosing.

== Cleaning the eggs ==

=== Outline and summary of the egg cleaning process ===

This stage consists, essentially, of cleaning the eggs using density separation, size separation, possibly a chemical
cleaning using the septic tank cleaner, Rid-x, and acid.

Here is my abbreviated outline and summary of the various stages involved in this whipworm egg cleaning protocol.<ref name="OvumVsEgg" group="note" />

# Once any cork with hookworm larvae has been separated off, I then have to prepare [http://veterinarymedicine.dvm360.com/recipe-sheathers-sucrose-solution Sheather's solution] <ref name="SheathersSolution" group="note" /> for a very rough initial density separation.
# I thoroughly mix the soil and stool into the Sheather’s in a stock pot with a dedicated strainer.
# A day or so later, I skim off the material from the top of the solution with a homemade hand scoop.
# I filter this material through a 30 micron mesh. <ref name="30MicronMesh1" group="note" /> This separates the ova, which pass though the mesh, from the stool debris, most of which remains on top of the mesh and can be discarded.
# I then filter the material using a 20 micron mesh. <ref name="WetMesh" group="note" /> <ref name="20-30Mesh" group="note" /> <ref name="PPip" group="note" /> <ref name="Sucking" group="note" /> This captures the ova, which remain on top of the mesh, while allowing most of the stool debris to pass through. Most estimates I’ve come across claim that whipworm eggs measure from about 58 to 50 microns long, and from 27 to 22 microns wide. <ref name="PPTPref" group="note" /> These two filtering steps in the procedure ideally mean that anything larger than 30 microns and smaller than 20 microns will have been filtered out.<ref name="TTeggs" group="note" /> So the material that's left at this point should have a reasonable concentration of eggs.
# I then centrifuge this material in a multistep procedure using both 1.130 and 1.200 specific gravity sugar water solutions. The eggs have a specific gravity between 1.130 and 1.200, and this allows their separation from the stool. <ref name="Buoyancy" group="note" /> <ref name="DensitySeparation" group="note" /> I then use a Pasteur pipette to suck the material off the top or bottom of the columns of liquid in the centrifuge tubes. This material is then re-filtered a second time, firstly using a 30 micron mesh to allow the eggs to pass through, and then using a 20 micron mesh to collect the eggs which remain on its surface. Later on, when I’m cleaning the centrifuge tubes, I find a [https://www.waterpik.co.uk/oral-health/products/dental-water-flosser/WP-450UK/ Waterpik] is very useful for cleaning the faecal material out of the bottom of them.

I don’t think it’s likely that [http://www.rid-x.com/ liquid Rid-x] harms the ova, <ref name="Ridx" group="note" /> but, because I’m not 100% certain about this, in the next step I divide my sample and, for now, follow both of these two separate procedures in parallel. <ref name="Ridx2" group="note" /> <ref name="cellulose" group="note" />
 
'''Then either:'''

<ol start="7">
<li>I put the eggs into a Rid-x septic tank cleaner and water solution for 4 weeks, changing the Rid-x and water every 7 days. This is done by pouring the week-old solution through 20 micron mesh, saving the material on top of it, putting it back through a 30 micron mesh into a [https://www.glad.com/food-storage/containers/mini-round/ Glad Mini Round container] that has about 8 small holes punched in the lid using the point of a pair of scissors, and then adding fresh Rid-x and water.<ref name="Glad" group="note" /> The Rid-x contains enzymes and bacteria that break down stool. The upshot is that the only visible material that will remain after 4 weeks of doing this will be the eggs and a few bits of (what I think is) cellulose.</li>
<li>After 4 weeks, I do the final 20 micron filtering, at which point I should see almost nothing but eggs under the microscope.<ref name="CleanCheck" group="note" /> I place the sample into a bath of mild 3.0 pH car battery (sulphuric/sulfuric) acid for three weeks. <ref name="AcidContact" group="note" /> Hopefully, this will kill any bacteria or protozoa that might have tagged along with the eggs, although there is no visible indication that this provides additional cleaning by removing any of the relatively larger contaminates that remain.</li>
<li>After three weeks, I neutralise the acid with baking soda, carry out a final 30/20/30 micron mesh filtration, and put the eggs into safe storage in water in a Glad container.</li>
</ol>

'''Or:'''

(The only difference here is that I've skipped the Rid-x part of the procedure.) 
As I said above, I doubt that Rid-x harms the eggs, but I’m not certain about this. So, for now, I keep a separate
batch of a couple of thousand eggs that haven’t been Rid-x treated in case my colony dies and I have to start over. If
I do have to use this sample to restart my colony, the sample is still clean enough such that I won't taste or smell
anything.

<ol start="7">
<li>I place the sample into a mild 3.0 pH car battery acid bath for three weeks.</li>
<li>After three weeks, I neutralise the acid with baking soda, do a final 30/20/30 micron mesh filtration, and put the eggs in safe storage into water in a Glad storage container.</li>
</ol>

=== Full description of the egg cleaning process ===

I assume that any cork bearing 3rd stage hookworm larvae will have already been removed from the culture,
although I still wear gloves and an apron during this procedure in case any stragglers remain. I carefully remove all
the cork, because, if any is left, it will float to the top of the Sheather’s solution later on and contaminate the
mixture.

<blockquote>

==== Preparing Sheather's Solution ====

To prepare Sheather's solution, ([http://veterinarymedicine.dvm360.com/recipe-sheathers-sucrose-solution], [http://www.jorvet.com/wp-content/uploads/2012/01/sheathers_sugar_flotation_solution.pdf]) I begin with 2.721 kg (6 lbs) of table sugar (sucrose) and 2,130 ml (72.02
U.S. fluid ounces) of distilled water. To measure out the sugar, I use a small 1kg (2 lb) capacity postal scale, and, to
measure out the water, I use a plastic graduated cylinder. (Plastic is better than glass because it's unbreakable.)<ref name="hydrometer1" group="note" />
I first sieve the sugar to remove any lumps. Then I heat the water in a 10 litre (11 quart) stock pot on the stove. I
don’t want to get the water too hot, since this might caramelise the sugar. After it’s beginning to get too hot to
comfortably put my fingers in, and well before it boils, I begin to gradually add sugar, using a small bowl to ladle it
from the larger container it’s in. I don’t just dump the sugar in, but shower it in, while stirring the mixture
constantly to avoid sugar build up and caramelisation on the bottom of the pot. Once all the sugar is dissolved, I
put the pot aside to cool. (If I want to speed up this part of the process, I sit the pot in cool water to reduce the
temperature more quickly.) Once the solution has cooled to room temperature, I use a large funnel to drain off
about ½ of it (about 2 litres [2 quarts]), for use during the later 1.200 and 1.130 centrifugings. I put this into 2
[http://www.rubbermaid.com/en-US/chug-bottles Chug Bottles] and store them in my refrigerator.

Next, I use a hydrometer to check the density of the solution, which should be close to 1.270, although this doesn't
need to be too exact at this stage.<ref name="hydrometer2" group="note" /> This can be done by putting some of the solution either in a Chug Bottle,
any other tall thin bottle, or by using my graduated cylinder.

==== The Sheather's Separation ====

Then, while wearing [http://www.amazon.com/Lehigh-Spontex-19005-Bluettes-Gloves/dp/B001B38WLY Bluettes] gloves, I use my hands to manually scoop the stool and soil from the container. I
then mix this by hand into the 1.270 Sheather’s solution in my stock pot using a large dedicated sieve.<ref name="ModSieve" group="note" /> The soil
and stool isn’t necessarily forced through the sieve, but worked against the interior side of the half sphere of the
sieve to work the lumps out as much as possible and get the solution as smooth as I can.<ref name="StoolCollect" group="note" /> At this point, as a
check, I take some of the stool-laced solution and check it for density. As long as its density is well over 2.200 the
separation procedure should be going according to plan.

I then let the mixture separate out for at least 24 hours, although I don’t actually know the separation rate of
whipworm eggs in Sheather’s solution. While doing this, I keep the lid of the pot slightly ajar, since the eggs might
not develop when stored with decaying matter such as faeces in a closed container.<ref name="JonesAWQuote" group="note" />

==== 30/20/30 Micron Mesh Filtering ====

Next, using my homemade hand-scoop, I skim off some solution from the top of the mixture.<ref name="HandScoop" group="note" /> Then I place the
material on top of a sheet of 30 micron mesh. This mesh is attached, by a wimpy rubber band, to the top of a
circular mouthed plastic party cup 9.5 cm (3¾ inches) in diameter across the top and 12.7 cm (5 inches) tall.<ref name="RubberBands" group="note" /> I
place the mesh over the cup, gently making a depression or pouch about 5 cm (2 inches) down into the mouth of
the cup with my curled fingers and/or fist because I want the pressure to be evenly distributed and not focused at
any one point on the mesh. I then put the rubber band over the cup, securing the mesh to it. The liquid solution is
then poured down into the depression in the mesh, to sit on top of it. I then use a plastic spoon, or the bulb of my
Pasteur pipette,<ref name="PasteurPipettesx2" group="note" /> and a spray bottle<ref name="Rubbermaid32oz" group="note" /> to gently work the material through the mesh as thoroughly as possible.
Since this requires so much water, I need to transfer the mesh several times from full cups to empty ones.<ref name="ThirdHand" group="note" /> <ref name="DechlorinatedWater" group="note" />
<ref name="30MicronMesh2" group="note" />
I’m eventually left with a remainder bolus of material that won’t go through. This will be later disinfected by
being boiled for 10 minutes, then allowed to cool, and finally discarded down the toilet.

Next, I take the mesh off the 30 micron mesh covered party cup(s), and take the liquid from all the cups and pour it
through clean 30 micron meshes a second time. Obviously, this time it should go much faster. This liquid will now
only contain material smaller than 30 microns.

Now I take the liquid and pour it through a 20 micron mesh, which is attached on top of another party cup in the
same way as the 30 micron mesh was. Because these 20 micron mesh covered party cups will typically fill up as I
repeatedly pour and spray the liquid through the 20 micron mesh, I'll often have to hand transfer the 20 micron
mesh with the liquid and material on top of it from a full cup to an empty one.<ref name="ThirdHand" group="note" /> I continue spraying this
residue, and changing and emptying the cups, until the water I’m getting in the cups is quite clear. Eventually I'll
have a small amount of residue on top of the 20 micron mesh that is reasonably clean. All of the eggs should be on
top of this mesh, and so I dispose of the liquid left in the cup(s) and later boil it.

I next use a mini pipette to suck up some of the material, and then put it on a slide<ref name="LampHeat" group="note" /> <ref name="MinimizeLoses" group="note" /> so that I can check with
my microscope to see if I have any eggs, although I have always had them in the past. Once I’ve confirmed the
presence of eggs, I suck the sample up with a Pasteur pipette that has the end cut off to widen its opening to better
suck up the material, and then expel it through a 30 micron mesh into a glad container.<ref name="Squirt" group="note" /> <ref name="PPip" group="note" /> Of course, this
container will also have some water in it.<ref name="Transferring" group="note" />

I then repeat this whole process of scooping from the 1.270 liquid and doing 30/20/30 micron mesh separation a
number of times until after a number of hours it looks as though I’ve gotten enough material. I then dispose of the
leftover material that remains in the stock pot by boiling it.

Then I take the 30/20/30 separated material and 30/20/30 filter it again as a further cleaning. This part should go
quickly.

==== Preparing 1.200 and 1.130 Sugar Water ====

Next, I need to prepare 1.200 sugar water from the Sheather's solution that is stored in one of the Chug bottles.<ref name="WarmChug" group="note" />
So I prepare my wet sponge hydrometer bed, and then take out my hydrometer.<ref name="hydrometer2" group="note" /> Then I measure the specific
gravity/ density of the solution by dipping the hydrometer into the solution in the bottle. It will be about 1.270 so I
will need to cut it with distilled water. I pour out a little of the liquid (perhaps 120 ml (0.5 cup), add distilled water,
then put the cap on and gently mix the solution by pointing the top of the bottle down and then back up several
times. I do this gently because I don't want foam to form in the liquid, since this can cling to the hydrometer and
make reading it more difficult. Once I've mixed the solution, I then open the bottle and, using my hydrometer,
measure the new density. As I'm getting close to my goal, I pour out and replace far less liquid to avoid
overshooting the mark. (If I did this, I would need to make 454 g (1 lb) worth of new 1.270 Sheather's, and then, in
the same way as before, sequentially add that to my solution to gradually increase its density.)<ref name="LongStorage" group="note" /> This process
takes a number of repetitions until I get a density of 1.200.

I then repeat this whole procedure with the other chug bottle, and make the 1.130 sugar water.

==== Centrifuging the Mixture ====

Next, I take my solution containing eggs and pour it onto <ref name="Transferring" group="note" /> a sheet of 20 micron mesh that is of course attached
to the top of a party cup. I then use a Pasteur pipette to put some of my 1.200 solution onto the 20 micron mesh
along with the egg-containing solution. The reason I'm doing this is because if I simply added this egg-containing
solution to a centrifuge tube there would be some chance that the water in it, with a specific gravity of about 1,
would change the resulting mixture in the tube when I add the 1.200 solution to it. I want to keep adding 1.200
solution to the egg-containing solution, letting it replace the 1 density liquid water as it's dripping through the 20
micron mesh, until the egg-containing solution has been largely transformed into a 1.200 solution. I keep on doing
this for, say, 5 minutes, by which time the fluid will have been replaced perhaps 6 times, which I'm guessing is
enough to get the egg-containing solution very close to 1.200. Then, I use a cut-off Pasteur pipette to suck up the
material on the mesh and squirt it into a centrifuge tube. I try to ensure that this liquid that is drawn off of the top
of the mesh will be about 1/5th of the total I’ll be putting into the tube, and the other 4/5ths will be 1.200 solution
that is drawn directly from the Chug bottle. This further ensures that the density of the solution in the tubes will be
correct. Also, when I fill a tube I only fill it to about 80% of its total height, leaving some margin for error to prevent
spills. Depending on how much material I have, I might fill from 1 to 6 tubes in this way. A centrifuge '''must be
absolutely balanced''', for safety, so, if I fill less than 6 tubes in this way, I will then fill the remaining tubes with the
1.200 solution from the Chug bottle, so that all the slots in my centrifuge are full.<ref name="UnbalancedCentrifuge" group="note" />

Once my centrifuge is fully loaded, I close it and let it run for 10 minutes at about 4,000 rpm, generating about
1,600 g-forces.<ref name="GForce" group="note" /> Since the eggs are lighter than 1.200, and almost all the other material mixed with them is
heavier than 1.200, the eggs should have risen to the top of the tube(s) by the time it stops, and almost all of the
rest of the stool should have sunk to the bottom. I now use a cut-off Pasteur pipette to suck approximately 0.5 cm
(0.2 inch) from the top of the column of liquid in each tube that had egg bearing material in it. When I'm sucking up
the liquid from a test tube with the pipette, I "tap" the pipette's mouth repeatedly vertically up and down onto the
top of the liquid, roaming/moving around and sucking the top of the liquid, looking for fresh material. I suck the
liquid up and then empty it onto a piece of 20 micron mesh fabric that's on a party cup. I then add more 1.200
solution from the Chug bottle to top off each tube, back to the 80% fill level that they had previously. Then I recentrifuge
the whole batch. The reason I repeat the centrifugation is that each time I pipette liquid from a tube I'm
only about 95% efficient at getting all the eggs, leaving some on the tube's upper sides, etc. So, if I do this three
times, I'll hopefully be getting virtually all of them.

The liquid I'm extracting from each tube needs to be kept constantly wet on the mesh by spraying it every once in a
while. When I'm done with the tubes for a round of centrifugation, I then vigorously and repeatedly spray the mesh
to both further clean the eggs and replace the 1.200 solution they are in with 1 density water. At this stage, I might
do an additional 30/20/30 micron mesh filtering to remove any relatively large, stray materials that might somehow
have been introduced.

I then repeat this whole procedure with the 1.130 solution, but of course now the eggs will be drawn up from the
bottom of the tubes using a non-cut-off Pasteur pipette.<ref name="DensitySeparation" group="note" />

==== Rid-x Treatment ====

Either: 
I put half my eggs into a Glad container with about a 1/3rd of a shot of filtered liquid Rid-x<ref name="FiltrationStep" group="note" />, and then fill the
container the rest of the way, about 75% full, with dechlorinated water.<ref name="WaterNote" group="note" /> Then I put the container away in a safe
place, in the dark, in my unplugged mini fridge. Every 7 days, for 4 weeks, I change the Rid-x by 20-30 micron
filtering this sample, and then adding fresh Rid-x and water. After 4 weeks I do a 30/20/30 micron mesh filtration.<ref name="ClosedSystem" group="note" />

'''Acid Treatment''' 
Next I put this Rid-x treated sample on a 20 micron mesh, drain the liquid, and then Pasteur pipette it into a Glad
container with as little water as possible. Then I pour in car battery acid, filling the Glad container about 75% of the
way.<ref name="Acid" group="note" /> Then I put it away in my unplugged mini fridge for 3 weeks. After three weeks, I take out the Glad
container and empty its contents into a 0.5 litre (16 fl oz) plastic container. I then add some more acid to the Glad
container to rinse it and get all the egg-containing acid solution out of and into the larger container. (NB: I do not
spray rinse the Glad container with water to clean it at this point because this would be adding water to acid.) I then
neutralise the acid by mixing it with baking soda, using a plastic spoon.<ref name="BakingSoda" group="note" /> I have to do this outdoors because the
fumes given off from this reaction are very caustic to breathe. Also I only add a little baking soda at a time to
prevent excessive foaming. Once no more foaming takes place when I add and stir in additional baking soda, and
there is also some free baking soda undissolved on the bottom of the container, the solution is then neutral pH = 7,
the same as water. I next 30/20/30 filter this solution to remove the baking soda powder, spray cleaning the eggs
thoroughly when they are on top of the 20 micron mesh, and then put the now finished eggs into safe storage in a
Glad container under water in my mini fridge.<ref name="AnotherWaterNote" group="note" /> I also attach a note made from an index card with the date and
contents.

T. trichiura eggs need to be kept well below 50ºC (122ºF), and well above 0ºC (32ºF), although there is some
disagreement in the literature about their ability to withstand cold. They should also be kept out of strong sunlight,
and, while they need oxygen to develop, they can probably tolerate low oxygen conditions. They will rapidly
succumb to desiccation, but if stored properly, it's possible for them to remain viable for more than five years.<ref name="EggsInStorage" group="note" />
They should not be in a closed container with decaying material.<ref name="QuotesFrom" group="note" />

Or: 
I treat the other half of my eggs in the same way as the paragraph above, except as I said they have not been treated
with Rid-x.

== Developing the eggs ==

I keep my eggs at around 27ºC (80ºF) for at least 4 weeks from the day of the bowel movement that they came from
before I dose. Of course, this includes their time spent in the culture, the acid and/or the Rid-x.<ref name="WhipwormEggsDevelop" group="note" />

== Dosing ==

When I’m going to dose, sources recommend that I don’t consume anything carbonated or hot for an hour both
before and after dosing. (I go further and only wash them down with dechlorinated water, and don’t consume
anything at all for an hour before and afterwards.) I start by getting the Glad container out of my mini fridge and
set up my microscope. I use a mini-pipette to draw up a small amount of liquid off the bottom of the container and
place a drop on a slide.

== Counting the eggs ==

I use 40 magnification to count the eggs. If the eggs are too clumped together to count easily I repeatedly suck them
in and out of the mini-pipette to break up the clusters. I can also accomplish this by doing a 30-20-30 separation. I
then start at the East or West side of the drop and count down vertically. I then move over, using the eggs at the
periphery of my vision as a marker for how far to move, and then repeat my vertical count. Once I’ve counted a
drop, I write the number down on an index card and, always keeping the slide horizontal, put it in my mouth like a
spoon and consume the drop. I repeat this process until I’ve dosed with the total number I want.

There doesn't appear to be a consensus on how long whipworms live.<ref name="WhipwormLiveFor" group="note" /> The estimates I've seen range from 1 to 8
years, and their lifespan might very well also depend on the biology of the human host. So this uncertainty means
that the dosing numbers and schedule will also be uncertain. In addition, somewhere between 10 to 20% of eggs
succeed in implanting, introducing still more uncertainty.<ref name="NoCitationForThis" group="note" />

I picked a rough midpoint estimate of a 3 year lifespan, and assumed that, on average, 15% would implant. So if I
dose at 400 every 6 months, this works out to 60 whipworms implanting per dose, and if they live 3 years on
average I will end up with 60 times the 6 doses in the 3 years before the first group in this series of them dies,
which equals a colony of 360 whipworms. The lower estimates of a 1 year lifespan and 10% implanting works out to
a colony of 80. The upper estimates of an 8 year lifespan and 20% implanting works out to a colony of 1,280. So, if I
assume my physiology is reasonably normal, the size of my colony might range anywhere from 80 to 1,280, with a
very rough midpoint estimate of 360.

</blockquote>
== Notes ==

<ref follow="cleanup" group="note">
Companies that sell these organisms carry out a far more rigorous cleaning process than I do, since I'm merely
self-inoculating. 
"But in short we use a combination of flotation, centrifugation, filtration, and flocculation to separate whipworm
from faeces, it is then incubated in an aerated one molar sulphuric acid solution for three weeks, then soaked in
Chlorhexidine for at least a day, then washed with various antibiotics and then suspended in a
buffered saline solution called M9."[https://groups.yahoo.com/neo/groups/helminthictherapy/conversations/messages/6527] 
Also, these companies’ donors have been thoroughly tested for contagious diseases, even though there's no
evidence that helminths are a vector for disease. (Meredith E. K. Calvert, Potential Transmission of Pathogens
Between Human Hosts by the Hookworm, Necator Americanus, 4 May 2008, available from the [https://groups.yahoo.com/neo/groups/helminthictherapy/files/ Files section] of the
'Yahoo Helminthic Therapy Forum'[https://groups.yahoo.com/neo/groups/helminthictherapy/info].)

</ref><ref follow="Bleach" group="note">While this will kill hookworm 3rd stage larvae, (See Note 1 in the [[NA_incubation:_very_detailed_method_by_Alana#Notes]].) I'm not certain how effective this is for whipworm eggs. Until I have more complete information I thoroughly clean everything and boil the liquid remains.

</ref><ref follow="Robust" group="note">I still get whipworm eggs even if my hookworm culture has failed completely. It seems that whipworms are more robust egg producers.

</ref><ref follow="TTeggs" group="note">
As discussed earlier, after culturing for 8 days it’s extremely unlikely that any viable hookworm eggs could still
be left, and, even if there were, the eggs of the hookworms, Necator americanus, average about 60 by 40 microns,
which means that they would be filtered out by the 30 micron mesh.
 
Also, it’s the width of whipworm eggs that is the relevant dimension for this filtration process, since the whipworm
eggs can be counted on to rotate appropriately when agitated in liquid on top of a 30 micron mesh surface such
that they will pass through it.

</ref><ref follow="OvumVsEgg" group="note">
An egg is the fertilised state of an ovum with the genetic materials of a male gamete. An ovum is the female
gamete. Since the nucleus of this reproductive cell contains only half the number of chromosomes that a usual cell
does, an ovum is a haploid cell. ([http://www.differencebetween.com/difference-between-ovum-and-vs-egg/ Difference Between Ovum and Egg].)

</ref><ref follow="SheathersSolution" group="note">
I do not use formaldehyde in my Sheather’s solution since this might harm the eggs.

</ref><ref follow="30MicronMesh1" group="note">
I bought my mesh locally, but I could also have obtained them through the supplier, Small Parts, that sells them
on Amazon:
* [http://www.amazon.com/Nylon-Opaque-Off-White-Length-microns/dp/B001D8GD4U/ref=sr_1_1?s=industrial&ie=UTF8&qid=1428015733&sr=1-1&keywords=30+micron+mesh Nylon 6/6 Woven Mesh Sheet, Opaque Off-White, 12" Width, 12" Length, 30 microns Mesh Size, 18% Open Area (Pack of 5)]
* and [http://www.amazon.com/Nylon-Opaque-Off-White-Length-microns/dp/B001D8ECDE/ref=sr_1_2?s=industrial&ie=UTF8&qid=1428015971&sr=1-2&keywords=20+micron+mesh++5 Nylon 6/6 Woven Mesh Sheet, Opaque Off-White, 12" Width, 12" Length, 20 microns Mesh Size, 14% Open Area (Pack of 5)].

</ref><ref follow="WetMesh" group="note">
To get the mesh filtration going, it can be helpful to wet the mesh, by spraying it, before starting.

</ref><ref follow="20-30Mesh" group="note">
It’s easy to get the 20 and 30 micron meshes confused. To be sure which I’m dealing with I check them with my
microscope. With experience I can also fairly well tell the difference by feel.

</ref><ref follow="PPip" group="note">
When the sample is on top of the 20 micron mesh I use a Pasteur pipette (which is cut off across about half an
inch from the tip to widen the opening) to suck the top of the mesh after the sample has been thoroughly rinsed. I
add water and then suck the mesh's surface repeatedly, pulling the open mouth of the pipette over the mesh gently
as I suck up the water. Even after all visible "dirt" of the sample material is gone there can be still a fair number of
eggs left behind on the mesh. I repeat this process a number of times past any visible remains to collect as much of
the sample as possible.

</ref><ref follow="Sucking" group="note">
When sucking the eggs off the top of the 20 micron mesh, fibres will often come off the mesh and get into the
sample. A simple way of removing them is to place a clean 30 micron mesh on top of the Glad storage container
when I'm putting the material into it. I expel the eggs and liquid onto the top of the mesh and then repeatedly spray
them through it into the Glad container, using dechlorinated water. This is why I often refer to my filtration
procedure as 30/20/30: first 30, then 20, and then 30 again. I have also taken to storing all my meshes in the cold
in my regular refrigerator (obviously not my unplugged mini fridge) in case there is some sort of deterioration going
on, although I have no idea if doing this is helpful. 
Although I haven't performed this experiment yet, another way of avoiding fibres from being sucked off the 20
micron mesh would be to simply remove the mesh from the cup and then flip it over on top of a 30 micron mesh
that is rubber banded on top of a party cup. Then I could spray through both meshes until the eggs had fallen off
the 20 micron mesh. Then remove the 20 micron mesh and spray any remaining eggs through the 30 micron mesh.
And finally transfer the contents of the cup into a Glad container. Of course, I would spray rinse the cup thoroughly
to ensure that all the eggs were transferred. So the 20 micron mesh shouldn't be producing fibres, and the 30
micron mesh should be removing them in any case.

</ref><ref follow="PPTPref" group="note">
[https://www.wadsworth.org/programs/ehs/pt Parasitology Proficiency Testing Program – Wadsworth Centre] / [http://www.bvsde.paho.org/bvsacd/cd65/SanitaryParasitology_Guide.pdf Integrated Guide to Sanitary Parsitology], p.12.

</ref><ref follow="Buoyancy" group="note">
[http://www.jstor.org/stable/3272719?seq=1%23page_scan_tab_contents The Buoyancy of Certain Nematode Eggs – Willi Sawitz, The Journal of Parasitology, 28(2), April 1942]. p 97. Table 2.

<div>
[[File:TTO Table2.png|link=]]
</div>

As an aside, the specific gravity of Hookworm Eggs is between 1.055 and 1.180. 
[http://www.jstor.org/stable/3272719?seq=1%23page_scan_tab_contents The Buoyancy of Certain Nematode Eggs, Willi Sawitz, Journal of Parasitology Vol. 28, No. 2 [Apr., 1942]]. p 101. 
It is also reported to be 1.055 in “The Specific Gravity of Hookworm Eggs” 
W. Sawitz, Am. J. of Tropical Medicine, s 1-19 (2), 1939, p 171-179 
Also, for comparison, here is the specific gravity of several other nematode eggs
* Toxicara canis Roundworm 1.09
* Ancylostoma Hookworm 1.0559
* Trichuris vulpis Whipworm 1.1453
So, if I'm using 1.200 separation, all the types of egg mentioned immediately above this sentence would float, and
all those mentioned immediately below would sink.
* Taenia Tapeworm 1.2251
* Physaloptera Stomach worm 1.2376
(Source: [http://www.jorvet.com/wp-content/uploads/2012/01/sheathers_sugar_flotation_solution.pdf Sheather’s Sugar Flotation Solution], Jorvet Jorgensen Laboratories, inc.)

</ref><ref follow="DensitySeparation" group="note">
I tightened up my density separation from 1.210, to both 1.200 and 1.130. With this different protocol I got a
fairly clean result, so I think that I will be using a 1.130 centrifuging along with the 1.200 one from now on. I now
suspect that faecal material changes its density over time. Previously a 1.210 separation, along with the 30-20 size
separation, gave OK results. This time, 1.200 and 30-20 resulted in a very dirty sample. Speculating here, if I had let
the faecal material age for more than a week I might have obtained better results. Since I hadn't done this, it was
only when I added the 1.130 separation step that I succeeded. Of course, when I do the 1.200, I'm pulling the eggs
from the top of the tube with a cut-off Pasteur pipette, since the eggs float in this density of solution. And, with the
1.130 density separation, I first suck off the top of the tube, and then discard this faecal-laden material. Then I refill
the top of the tube with 1.130 solution, re-centrifuge the tubes, and then again suck off the top of the tubes and
discard this material. Then I pull the eggs up from the bottom of the tubes using a non-cut-off Pasteur pipette.

</ref><ref follow="Ridx" group="note">
I only use liquid, and not powdered, Rid-x because the powdered form will introduce particles into the final
result.

</ref><ref follow="Ridx2" group="note">
If I'm still producing whipworm eggs after a number of years of dosing with Rid-x treated eggs then I’ll be
confident that the Rid-x doesn't harm them. The quickest way to be sure about this would be for me to prepare
some eggs using the protocol that includes Rid-x and then see if an inoculated person (obviously, someone who
didn't have any whipworms before this) produced eggs after about three months, since the adults mature in about 3
months. (Marcial-Rojas, RA. 1971. Pathology of Protozoal and Helminthic Diseases with Clinical Correlation.
Baltimore: Williams & Wilkins. p. 661.) Unfortunately, I can't address this question because I have no close family
members to perform this experiment with.

</ref><ref follow="cellulose" group="note">
I tried removing the last bits of what I think is cellulose using cellulase enzyme, but this destroyed the eggs.
Cellulase enzyme comes in a variety of forms, so for all I know some other form might not have harmed the eggs.
But cellulase is hard to obtain, there is very little of any contamination left at this point and what is left is probably
cellulose, as found in vegetables (celery, for example, [http://www.nytimes.com/2006/07/25/health/25real.html?_r=1& is mostly cellulose]); and, since I had run out of patience at
this point, I abandoned this line of enquiry.

</ref><ref follow="Glad" group="note">
I punched small holes in the lids of all the Glad containers used in this protocol because the eggs need some
oxygen, and they should also not be held in a closed container with possibly decaying material. I'm now storing my
Glad container, which contains my whipworm eggs, within an open topped square plastic container. So the Glad
container sits in the middle of a nest of surrounding aluminium foil, which serves as a buffering cocoon, all inside
the larger square container (technically a cube). This arrangement reduces the possibility that I might accidentally
overturn the Glad container, and so lose my whipworm eggs. Of course, this arrangement is kept inside my minifridge.

</ref><ref follow="CleanCheck" group="note">
When I’m examining how clean the eggs' are, I use a two-step process.
# I first focus on the plane of view at the eggs' level/depth, scanning for debris within the eggs.
# I then focus up to the surface plane of the water bubble that is on the slide, scanning for any floating debris

There are some bits of stuff (probably cellulose) left, but I remind myself that other food I eat has undesirable
things in it too. Popcorn is a special favourite. “1 or more rodent excreta pellets are found in 1 or more subsamples,
and 1 or more rodent hairs are found in 2 or more other subsamples” ([http://www.fda.gov/Food/GuidanceRegulation/GuidanceDocumentsRegulatoryInformation/SanitationTransportation/ucm056174.htm Defect Levels Handbook])
 
 
Also, I remember hearing a radio program that reported that about 80% of the food ladles/handles used in
smorgasbords have urine and faecal material on them because people didn't wash their hands after using the
bathroom. The inside surfaces of a public bathroom are very likely worse. ([http://serendip.brynmawr.edu/exchange/node/1839 Think before you flush or brush]) See
also, [http://wfncnews.com/3278/close-toilet-lid-flushing-brush-poop Close the Toilet Lid When Flushing or Brush With Poop. Your Choice] and [http://msgboard.snopes.com/cgi-bin/ultimatebb.cgi?ubb=get_topic;f=35;t=001754;p=0 Topic: Close the lid before you flush?]
 
 
I also remind myself that faecal transplants (using either the oral or anal routes) are becoming more acceptable to
allopathic medicine. And the way our body’s [http://en.wikipedia.org/wiki/Gut_flora#Carbohydrate_fermentation_and_absorption carbohydrate fermentation and absorption] works means that a
significant part of our food is pre-digested by our gut bacteria. In other words, we are eating their faecal materials.
As if I needed any more gross facts to get over any squeamishness over contamination, I remind myself that
[http://www.quirkyscience.com/alcohol-is-yeast-urine/ alcohol is yeast urine], that [http://news.hjnews.com/cache_magazine/don-t-be-scared-off-by-the-unflattering-process-of/article_1b56f7b2-618a-11e2-97b0-001a4bcf887a.html cheese is bacteria faeces], that data has confirmed the transmission of fecal coliforms in
communal bathrooms at a university, that [http://www.sciencedaily.com/releases/2015/06/150602130650.htm toothbrushes can serve as a vector for transmission of potentially pathogenic organisms], and that
[http://news.yahoo.com/theres-probably-poo-ground-beef-care-212130887.html?soc_src=mail&soc_trk=ma there's probably poo in our ground beef].

</ref><ref follow="AcidContact" group="note">
I have to make sure that micron mesh doesn't come into contact with the acid, or [http://www.chemguide.co.uk/organicprops/amides/polyamides.html it will dissolve].

</ref><ref follow="hydrometer1" group="note">
Although 500 ml size might be better for measuring the distilled water for the Sheathers solution, I think that
250 ml is a convenient size for using a hydrometer to check the solution’s density

</ref><ref follow="hydrometer2" group="note">
I must be very careful when handling a hydrometer, since they are extremely fragile. When I take it out of its
box I try to always hold it with both hands, one on the stem and one cupped underneath it. To make a soft bed for
it when I’m putting it down on the table I lay out three rectangular wet sponges 7.5 cm x 11.5 cm x 1.25 cm (3 x 4½
x ½ inch) in a 34 cm (13½ inches) long line on the table. When I lower it down into any solution I’m careful to not
even allow it to bang slightly against the sides or bottom of the container. Then I carefully rinse it under flowing
water in the sink and put it away as soon as I’m done using it.

</ref><ref follow="ModSieve" group="note">
I modified a sieve by cutting the handle off with a hacksaw, and then used [http://www.amazon.com/Shoegoo-110231-Shoe-Adhesive-Clear-1-Fl-oz/dp/B008AF1NG2 Shoe Goo] to cover over the resulting sharp edges.

</ref><ref follow="StoolCollect" group="note">
It’s almost essential to have started with soft stool. If the stool is hard to begin with, this part of the process
will be many times more difficult, and could take hours. So I eat a large quantity of celery the day before starting
the culture to try to ensure that it will be soft. However, this means that when I have to defecate, I have to do so
immediately. Since I will typically be out and about, I need to carry a couple of suitably sized Ziploc type bags to
double bag the stool in, and then quickly get it back home and into a culture. I have considered using a dedicated
blender to emulsify the stool, but haven't yet done this experiment, and I don't know whether this might damage
the eggs.

</ref><ref follow="JonesAWQuote" group="note">
“Moderate drying inhibits embryonation; heat from the sun’s rays, freezing temperatures, decaying matter such
as stored faeces, various strong chemicals, and prolonged drying kill the eggs and embryos.” (Jones AW.
Introduction to Parasitology. 1967. p 221)

</ref><ref follow="HandScoop" group="note">
I made my hand-scoop from a soft plastic cereal bowl. I cut the plastic with kitchen scissors to fashion a 10 cm (4 inches) wide and 2.5 cm (1 inch) deep scoop.

</ref><ref follow="RubberBands" group="note">
I use the weakest rubber bands that will still do the job, since I don’t want to take the chance of tearing the
mesh. Also, in this paper, when I'm referring to a "cup" I'm almost always referring to the party cups listed in the
materials section.

</ref><ref follow="PasteurPipettesx2" group="note">
I ended up with two sizes of Pasteur pipettes, one about 15 cm (6 inches) and the other about 22 cm (8¾
inches). The 6 inch ones are the ones I use most frequently. The 22 cm (8¾ inches) ones, which are typically used
for perfume applications, probably aren't needed, but they have proved useful for sucking up material from deep
inside a bottle of the 1.130 or 1.200 solutions when they’re getting empty. If my test tubes were much deeper, the
22 cm (8¾ inches) pipettes also would be needed if I ever needed to draw up material from the bottom of a tube.

</ref><ref follow="Rubbermaid32oz" group="note">
I particularly like the [http://www.amazon.co.uk/Rubbermaid-Plus-Heavy-Spray-Bottle-FG6C13PR06/dp/B00DLJ0A0C/ref=sr_1_58?ie=UTF8&qid=1436047997&sr=8-58&keywords=heavy+duty+spray+bottles Rubbermaid 32oz Heavy-Duty Spray Bottle] because it has a strong spray, I find that I
need to spray down at a somewhat horizontal angle into the mesh in the party cup, but, given the way the bottle's
internal mechanism works, the sprayer stops working as the bottle empties, and, even if it’s still half full, it won't
spray. Due to this, I find it's best to have a number of spray bottles, otherwise I need to refill them frequently.
About six appears to be the ideal number.

</ref><ref follow="ThirdHand" group="note">
Changing micron mesh to a new party cup when it has egg bearing material on top of it is a little tricky, since I
have to get the rubber band off, transfer the mesh to a new cup, and then get the rubber-band back on, all the while
keeping the mesh depressed so that I don’t spill the material that is sitting on top of it. I don’t want the depression
in the mesh to become flat. This is really a two person job, and the ideal one-man solution would probably involve
some sort of spring loaded device that snapped down around the mesh and held it on the container, but here’s my
method for one person doing it while also utilising a rubber band to keep the mesh in place. To ease the task, I
created a simple third hand by cutting off the top half of a 0.7 litre (24 Am fl oz) (upside down [http://mathworld.wolfram.com/ConicalFrustum.html conical frustum]-shaped)
[http://56bellps.global2.vic.edu.au/files/2014/02/Tub-q9ntzb.jpg yogurt container]. I had to make sure that the cut edges were smooth with no [https://en.wikipedia.org/wiki/Burr_%28edge%29 burrs] so that they won’t later
snag the mesh. The resulting half yogurt container's dimensions are about 6.3 cm (2½ inches) tall, with a 9.5 cm (3¾
inches) diameter circular mouth, and an 8.25 cm (3¼ inches) diameter base. The party cups are about 12.7 cm (5
inches) tall, with a 9.5 cm (3¾ inches) diameter circular mouth, and a 5.7 cm (2¼ inches) diameter base. This
arrangement leaves about a 1.25 cm (½ inch) circumference gap between the two when the cup is placed down into
the container. So I place the cup, with the mesh rubber-banded onto it, down into the container, tucking the mesh
fabric down into the gap space that's between them. When I want to transfer the mesh, I hold the top of the mesh
in place, cupping one hand down onto it, grasping the top rim of the cup gently but firmly through the mesh. Then,
with the other hand, I pull the rubber band away from the side of the mesh on the cup and slip the band out and
down onto the outer container, transferring the band to it. Then I lift the cup and set it next to another fresh and
empty cup. Next I lift up the mesh and transfer it to the new cup. I then put this down into the half yogurt
container, again tucking in the mesh. Then, while holding the top mesh in place as before, I slip the rubber band off
the half yogurt container, and up and onto the new inner cup, such that the rubber band is now holding the fabric
in place once again. If I do have to use my gloved fingers to adjust and force the mesh back down a bit into the new
cup, increasing the depression, I then spray my fingers over the mesh so any eggs that might be clinging to them
won’t be lost, but instead will fall into the mesh.

</ref><ref follow="DechlorinatedWater" group="note">
If I’m doing a lot of scoops of material this will take a lot of water, so, to have enough dechlorinated water, I fill
up two 15 litre (16 quart) containers. I use dechlorinated water for this work, although I’m not completely sure that
this is necessary. Dechlorinated water is easily prepared by filling a plastic container, 15 litre (4 US gallons) size
with cold tap water, and then letting it stand with the lid off for 24 hours. (I always use cold water because there is
less risk of contamination. Hot water can much more easily leach contaminates from pipes.) Tiny bubbles form on
its inside surfaces, which I remove by tapping the container’s sides, and then scraping the remaining ones off using
a kitchen knife. Then I snap on its sealing top, and use the water as needed. I don’t use distilled water, since I don’t
know if the eggs can tolerate the osmotic problems that might be associated with this, and I'm guessing that
normally mineralised water is what they are most comfortable in. There are jug water filters ([http://www.brita.co.uk/brita/pages/product/list.grid?subshop=cpd_shop&lang=en&filter=cpd_shop_pitcher&country=GB Brita Jug Filters]) and
filters that attach to the home tap/faucet (
* [http://www.amazon.co.uk/Kitchen-Simple-Faucet-Filter-Purifier/dp/B00CM6RQBG/ref=sr_1_4?ie=UTF8&qid=1440276846&sr=8-4&keywords=tap+mounted+filter Home Kitchen Simple Easy Faucet Tap Water Clean Filter Double Purifier Head Pp]
* [http://www.amazon.co.uk/Culligan-FM-15A-Advanced-Faucet-Filter/dp/B00PRN88TK Culligan FM-15A Advanced Faucet Filter Kit]
** see [http://culligandiy.com/Faucet-Filters Culligan Compact Faucet Mount Filter Installation and Operation Instructions]
and some of these claim to remove over 97% of the chlorine, but I haven’t tried any of these to see whether they might work.
 
Many areas use chloramine instead of free chlorine to disinfect drinking water. If this were my situation, I would
first carry out a test to establish whether this had any adverse effect on the eggs. If it did, then I would either have
to investigate using one of the products designed for tropical fish owners, e.g., [http://eveliens.hubpages.com/hub/Aquatics-Product-Review-Seachems-Prime Seachem Prime],
[http://www.aquahobby.com/products/e_amquel_plus.php AmQuel] or [http://www.amazon.co.uk/gp/product/B00027ZTCU?*Version*=1&*entries*=0 Ammo-Lock],
([http://www.thetropicaltank.co.uk/rev-cond.htm this site presents a list of this type of product]), investigate using bottled water, distilled water, or water
from a local natural source; or even, as a last resort, consider installing an expensive and complicated multistage
filter system.
* [http://www.thetropicaltank.co.uk/chlor.htm Chlorine and Chloramine in the aquarium - The Tropical Tank]
* [http://www.chloramine.org/chloraminefacts.htm Chloramine Facts - Citizens Concerned About Chloramine (CCAC)]
* [https://www.cityoftulsa.org/city-services/water/chloramine-information/chloramine-faq.aspx Frequently Asked Questions - Chloramines - The City of Tulsa]
* [http://en.wikipedia.org/wiki/Chloramine#Removing_chloramines_from_water Removing chloramines from water - Wikipedia]

The only really practical way to remove chloramine is to use a water conditioning product such as those mentioned
above, or any one of numerous other brands. Only a couple of chemicals are used for aquarium dechloramination,
and all the brands on the market use one or the other of these. Adding the prescribed amount to the water (usually,
one teaspoon of conditioner to 38 litres [10 US gallons] of water) more or less instantly renders this safe for aquatic
animals ([http://answers.yahoo.com/question/index?qid=20120227201829AAQikQR How to remove chloramine from tap water?] and [http://www.thetropicaltank.co.uk/rev-cond.htm Review- Water Conditioners and Dechlorinators]).

</ref><ref follow="30MicronMesh2" group="note">
It takes about a litre (quart) of water per scoop of egg-containing solution to work it thoroughly through a 30
micron mesh. This can take a while, and so I find it useful to have several 30 micron meshes with party cups going
at once. Since I am first going from 1.270 separated stool, it’s likely that, when using a 30 micron mesh, this mesh
will become clogged with sediment. If a mesh does becomes clogged I spoon transfer the contents to another 30
micron mesh, and then heavily spray as much of the remaining material as I can through the mesh. Then I remove
and clean the mesh.
 
The meshes need to be hand cleaned with warm soapy water and rinsed thoroughly after, and between, each time
they are used. If I’m using them again right away I also then spray rinse them with dechlorinated water, although I
doubt this is necessary. If I’m drying them, I hang them with a clip freely suspended in the air, like
[https://repetitivemeasures.files.wordpress.com/2012/04/20120407_171701.jpg photos drying in a dark room] so that the wet meshes do not rest on anything from which they might accumulate contaminants. I
would only want to use clips similar or identical to the magnetic clips listed in the Materials section rather than
strong metal clips, such as [http://www.amazon.co.uk/Binder-Clips-Assorted-Silver-Package/dp/B00006IBA2/ref=sr_1_1?ie=UTF8&qid=1438112141&sr=8-1&keywords=Silver+Binder+Clips Binder Clips],
since the area of contact between the jaws of the clip and the mesh needs
to be as large as possible to more evenly distribute the pressure on the mesh. A clip that grips in a way that’s more
like two fingers pressed together is better than one that acts like a knife edge pressed against a hard surface. A
hand held blow dryer will dry a micro-mesh very quickly, although '''I have to make sure I don’t get it too hot'''.
 
When cleaning Pasteur pipettes with washing-up liquid/dish soap and water, I alternately suck and squirt some of
the soapy water several times. Then I do the same with clean water. When rinsing them out I find it sometimes
useful to point them up into the stream from the tap and repeatedly fill them and then squirt out the liquid rapidly
upwards. The 22 cm (8¾ inch) ones can be very difficult to rinse the soap out of.

</ref><ref follow="LampHeat" group="note">
I have to be careful to not leave a drop on a slide unattended too long, since the heat from my light will
evaporate it over time.

</ref><ref follow="MinimizeLoses" group="note">
I’m aware that each time I handle the eggs I risk losing some, so the fewer times I handle them the better. In
any case, to minimize the loses, I rinse my Pasteur pipettes after I've just finished sucking up and expelling a
sample so as to not leave any eggs that might be clinging to the pipette’s inside or outside. I do this by drawing up
fresh dechlorinated water from a shot glass and expelling it into the mesh.

</ref><ref follow="Squirt" group="note">
I never squirt any liquid from a Pasteur pipette back into the Chug bottle because there’s too much risk of
contaminating the solution. Instead, I dispose of it, even if I know it’s completely clean. In the case of any larger
debris appearing, this can be removed by putting the Chug bottle liquid through a 20 micron mesh.

</ref><ref follow="Transferring" group="note">
Later on, when I might be transferring a solution with eggs in it from any container, I spray up into the
container onto its internal bottom and internal sides a number of times, repeatedly letting the liquid empty. Most of
the eggs will typically be on the bottom of the container since they sink in water. When I’m transferring eggs from a
microscope slide back onto a mesh I pick up the slide, hold it over the mesh and spray it a number of times, letting
the liquid drip off the slide and onto the mesh.

</ref><ref follow="WarmChug" group="note">
I like to get the Chug bottles out and warm them up a little, although I very much doubt this makes any
difference.

</ref><ref follow="LongStorage" group="note">
I’ve found that storing the 1.130 and 1.200 solutions in the refrigerator for longer than a few weeks isn’t worth
the trouble. Even in a sealed container, they develop “fibres” in them, which I think are condensed sugar. These can
be filtered out with a 20 micron mesh, but then the density can also change slightly, and might need to be adjusted.
I find it just as easy to simply make 2 new chug bottles’ worth, 2 litres (2 quarts) of it each time.

</ref><ref follow="UnbalancedCentrifuge" group="note">
An unbalanced centrifuge could easily become unstable, and therefore dangerous.

</ref><ref follow="GForce" group="note">
I don't know the eggs' tolerance level for g-forces, but I'm assuming it's reasonably robust. I’ve read of a
specialised method for separating ova using Eppendorf tubes at 13,000 RPM, however I don’t have that citation.
Other technical sources I’ve consulted typically talk in terms of units of 2,000 RPM and 500 g-forces. (Bowman DD.
Georgis’ parasitology for veterinarians, 7th ed. Philadelphia: WB Saunders. 1999. p 287.) Potentially I could address
the question of whether or not 1,600 g-forces harm the ova in the same way as I described in note 16.

</ref><ref follow="FiltrationStep" group="note">
This filtration step is very likely not necessary. Liquid Rid-x easily pours through the 20 micron mesh with no
apparent residue, except for foam created by shaking the bottle. (The instructions on the Rid-x bottle say to shake
the bottle, so I do this each time I use some.)

</ref><ref follow="WaterNote" group="note">
I'm only using either distilled or dechlorinated water throughout this whole protocol.

</ref><ref follow="ClosedSystem" group="note">
A closed system would become contaminated with breakdown products and toxins. So I need to periodically
filter the sample to remove them, and then introduce fresh materials. Also the pH could shift, which could slow or
stop growth. Four weeks might be overkill, but I'm being thorough here. Extending the Rid-x treatment beyond 4
weeks does not clean the sample any better.

</ref><ref follow="Acid" group="note">
I always, and only, add acid to water, [http://antoine.frostburg.edu/chem/senese/101/safety/faq/always-add-acid.shtml never water to acid]. When acid and water mix they create an exothermic
reaction, which means the reaction releases heat. If I were to add water to acid this could conceivably boil and
splatter the mixture, which could be potentially dangerous, although, since I'm using a mild acid, this possibly isn't
extremely dangerous, but I'm following correct lab practice at all times in all of this. And if I ever did use a stronger
acid then it indeed would be very dangerous to have that acid splatter. I don't want to get into bad habits that might
prove disastrous in the future.
 
Even though my Ph meter seems flaky, (perhaps all of them are) and will jump around strangely, it eventually
settles down such that the results make sense and I believe them. It indicates that the car battery acid is stable and
doesn’t appear to change pH even over a long period of time. According to the meter, when it’s been calibrated, the
acid I have in its original container is still a pH of 3 after a year and a half.
 
The pH scale runs from 0 to 14, with 7 being neutral. So from 0 to 6.9 is the acidic range, and from 7.1 to 14 is the
basic range. The acid bath with the eggs in it also remains a pH of 3.0 for the three weeks the eggs are soaking.
* pH = 0 => Battery Acid which is very strong Hydroflouric Acid
* pH = 1 => Hydrochloric Acid secreted by stomach lining
* pH = 2 => Lemon Juice, Gastric Acid, Vinegar
* pH = 3 => Grapefruit Juice, Orange Juice, Soda
* pH = 4 => Tomato Juice, Acid Rain
* pH = 5 => Soft Drinking Water, Black Coffee
* pH = 6 => Urine, Saliva
* pH = 7 => "Pure" Water
* pH = 8 => Sea Water
* pH = 9 => Baking Soda
* pH = 10 => Great Salt Lake, Milk of Magnesia
* pH = 11 => Ammonia Solution
* pH = 12 => Soapy Water
* pH = 13 => Bleach, Oven Cleaner
* pH = 14 => Liquid Drain Cleaner such as "Draino"
When I’ve looked at the pH that providers and laboratories seem to be using it seems that the ideal pH should be
between 0 and 2, inclusive, e.g.

* “... a composition for storage and development of eggs from helminthic parasites, where the composition further comprises a liquid carrier having a pH value of below 7 at a temperature of from 10 C to ambient temperature. The liquid carrier can be sulphuric acid, H2SO4 with a pH in the range of from 0 to 2, ...” ([http://www.freepatentsonline.com/y2010/0124574.html Link])

* “… it is then incubated in an one molar sulphuric acid solution for three weeks”…it is then incubated in an aerated one molar sulphuric acid solution for three weeks, then soaked in Chlorhexidine for at least a day, then washed with various antibiotics and then suspended in a buffered saline solution called M9.” ([https://groups.yahoo.com/neo/groups/helminthictherapy/conversations/messages/6527 Link])

* “Trichuris infective.- Trichuris spp. infective egg, was incubated for a period of four weeks at 26 ºC (78.8 F) “in a solution of 0.1 N” (1 pH) “sulfuric acid, obtaining the development to a stage 2 larva (L2).” ([http://ruby.fgcu.edu/courses/davidb/50249/web/tt6.htm Link])

Note: Talking about Normal acid strength can be confusing, since people do not always distinguish between dissociated sulphuric acid in water, versus non-dissociated pure sulphuric acid not mixed in water. There is a difference in strength of a factor of 2 between these.

* “0.12N is indeed 0.12 normal. In short, there are 0.12 moles of hydrogen ions per liter. Since this is sulfuric acid, it is 0.06 moles of” (non-dissociated) “sulfuric acid per liter.” ([http://chemistry.stackexchange.com/questions/19303/what-does-0-12n-mean-on-a-bottle-of-sulfuric-acid Link])

So these three sources have given pH numbers for soaking eggs as ranging from either 0 to 2, or 0 to 1.
 
When working with acids it is often necessary to convert from molarity to pH. For this immediate discussion I'm
only concerned about sulphuric acid, since how the acid dissociates in water makes a difference. 1 mole of nondissociated
pure H2SO4 that's then put in water, such that it results in 1 litre of total solution, will dissociate into 2
moles of H- ionic atoms and 1 mole of SO4+ ions. Since I get 2 hydrogen ionic atoms and 1 SO4 ion from each
H2SO4 molecule this will result in a 2 molar acid from one mole of H2SO4, since the number of H- ion atoms are
what determine the strength of the acid. The pH is defined as pH = -log (the molar strength), which equals the (base
10) positive log of (1/molar strength). So for a molar strength of 1, this means the pH will be log (1/1) = 0.
 
Dissociated Sulphuric Acid in water ([http://www.miniwebtool.com/log-base-10-calculator/ Log Base 10 Calculator])
{|
|'''Molar Strength,'''
|'''pH'''
|-
|1
| 0.0
|-
|½
| +0.3
|-
|¼
| +0.6
|-
|1/8
| +0.9
|-
|1/10
| +1.0
|-
|1/100
| +2.0
|-
|1/1,000
| +3.0
|}

I confirmed these results using this [http://www.sensorex.com/support/more/ph_calculator on-line pH Calculator]. For each calculation I had to enter 4 pieces of information. For example,
# The acid was “sulfuric,” since another acid might produce a different number of H+ ions per mole from the original non-dissociated molecule. Under “Volume Method”
# The concentration of acid, 1.0 moles
# The volume of acid 1.0 liters
# The total volume of the final solution 1.0 liters.

This gives a pH of 0.
 
I also confirmed these results at: [http://www.ochemonline.com/Acid_and_Base_Concentrations Approximate pH for different concentrations of various substances]
 
From the above table it’s clear that the pH scale is based on powers of 10. That is, a 3.0 pH acid is 100 times weaker
than a 1.0 pH acid, and it is 1,000 times weaker than a 0 pH acid. So, if I’m using sulphuric acid and ideally want the
pH to be from 0 to 2, then my molar strength should be from 1 to 1/100 of a mole of H+ ions per litre. Which
means the amount of the original H2SO4 non-dissociated molecules should be from ½ of a mole to 1/50th of a mole
of them. If I pick a mid-point goal of a pH of 1, since this equals the strength of stomach lining acid, then I would
want a 1/10 molar final solution, or 1/20th of a mole of pure H2SO4 that was then dissociated into water, such that
it added up to one litre of total final solution.
 
I considered making my battery acid stronger by boiling it, but this seems dangerous.
([http://chemistry.about.com/od/makechemicalsyourself/a/How-To-Make-Sulfuric-Acid-At-Home.htm How to Make Sulfuric Acid at Home].)
Also, the reduction necessary to go from a pH of 3 to 1 would be a factor of 100. So I'd have to buy 100
bottles of battery acid, at a cost of about £640 ($1,000/€895) and then reduce it to 1 bottle. The alternatives of
lemon juice or vinegar, both with a reported pH of 2 (see the above table), didn't seem attractive because these
might introduce additional biological material contaminates. Also, lemon juice doesn’t seem to be a good enough
disinfectant. ([http://www.aarp.org/home-garden/housing/info-03-2010/myth_buster_can_you_sanitize_kitchen_tools_with_lemon_juice_and_salt_.html Myth Buster: Can You Sanitize Kitchen Tools With Lemon Juice and Salt?]) White vinegar is reported
to be a reasonable disinfectant: “white vinegar is very effective at killing E. coli, salmonella and shigella
bacteria,” ([http://www.aarp.org/home-garden/home-improvement/info-03-2011/natural-cleaning-products-do-not-work.html Do 'Green' Cleaners Work?]) But when I tested the pH of the white vinegar I had, it came out as about 6.0,
which was worthless. [http://www.flinnsci.com/store/Scripts/prodView.asp?idproduct=21284 Flinn Scientific] sells 1 pH = 0.1 molar sulphuric acid, as does the [http://www.ebay.com/itm/SEOH-Sulfuric-Acid-0-1-Molar-Astm-D-3712-1-1L-/111628797746?pt=LH_DefaultDomain_0&hash=item19fd984732 Seoh Corporation]. However,
such scientific supply companies typically won't sell such products to individuals, but only to schools and similar
institutions. Having said all this, I have never become ill from my dosings when I’m using 3.0 pH battery acid to kill
any bacteria and protozoa that might remain in my sample.

</ref><ref follow="BakingSoda" group="note">
I’m careful to only add a little baking soda at a time, since the reaction generates a lot of foam. I also try to
make sure the plastic spoon is non-BPA, types 1, 2, 4, or 5.

</ref><ref follow="AnotherWaterNote" group="note">
If it becomes necessary to add more water because of evaporation over time, I add distilled water to the Glad
container. Using distilled water keeps the mineral concentration from gradually building up. If I thought it was
necessary I could filter and change the water with dechlorinated water every year or so. However, whipworm eggs
are capable of developing while in salt water, so they must be very tolerant of such mineral build-up.
 
(I’m assuming that in the following article the term “trichurids” includes Trichuris trichiura.)
 
“The eggs of the ascarids and trichurids attained the mobile larvae stage in 91-94% of the cases by the 25th day. It
was thus proved that clean sea water does not prevent development of geohelminth eggs within the optimum range
of temperature.” 
(Akademiia nauk SSSR. Contributions to helminthology: published to commemorate the 75th birthday of K.I.
Skrjabin. 1966. Verusalem: Israel Program for Scientific Translations. p. 802.)

</ref><ref follow="EggsInStorage" group="note">
'''The life expectancy of the eggs in storage.''' 
“... Likewise from the experiments of Davaine (1863), Morris (1911) and Epstein (1892) in which the eggs of Ascaris
and Trichuris remained infective as long as five years in preserved faeces it has been argued that eggs of these
species probably remain viable and infective in nature a corresponding length of time. This opinion has been
supported further by the results of experiments by Baillet (1866) and Cram (1924) who found Ascaris and Trichuris
eggs to be very resistant to freezing temperatures.” 
(Harold W. Brown, [http://www.jstor.org/stable/3271397?seq=1%23page_scan_tab_contents Studies on the Rate of Development and Viability of the Eggs of Ascaris lumbricoides and Trichuris trichiura under Field Conditions]. The Journal of Parasitology. 1927 Sep;14(1):1-15) 
"These eggs may remain viable outside of the host for many months if they lie in moist areas, but development
of the larvae within them may be delayed by dryness or cold, and when a favourable weather change occurs, the
bulk of the accumulated organisms continues development, thus making possible a massive infection."
(Noble ER, Noble GA. Parasitology: the biology of animal parasites. Philadelphia: Lea & Febiger. 1964. pp 373-374)

</ref><ref follow="QuotesFrom" group="note">
The quotations below are all from, [http://www.bvsde.paho.org/bvsacd/cd65/SanitaryParasitology_Guide.pdf Integrated Guide to Sanitary Parasitology]. (World Health Organization,
Regional Office for the Eastern Mediterranean, Regional Centre for Environmental Health Activities, Amman – Jordan. 2004. pp 25, 29-31, 36.)

* "A comprehensive resume of the time-temperature requirements for the destruction of Ascaris, Trichuris, Taenia and hookworm eggs is given by Feachem et al. (1983). In general, temperatures above 60°C (140ºF) are rapidly lethal to eggs." p. 25.

* "Exposure of eggs and partly developed embryos to freezing temperatures (-9°C to -12°C) resulted in a high percentage of fatality in Trichuris but had no apparent effect on Ascaris. It was also found that the further Trichuris eggs are developed, the less resistant they were to freezing" p. 27 (Nolf, 1932)

* "Trichuris eggs were much more resistant to the effects of the light. The difference in susceptibility was not definitely understood, but it was suggested that the dark pigmentation of the outer covering of the Trichuris eggs probably offers them considerable protection from the shorter light rays. It was also demonstrated that a very short exposure was sufficient to prevent a large percentage of Ascaris eggs from reaching embryonation, and slightly longer exposure was completely lethal to them" p. 29 (Nolf, 1932)

* "Trichuris eggs were killed at slightly lower temperatures (52°C to 54°C) for a shorter time than were Ascaris eggs (Nolf, 1932). The additional minutes exposure plus one degree rise in temperature reduced the percentage of surviving Trichuris eggs from 67% to 14% and the Ascaris eggs from 93% to 26%. Further, if the eggs of Ascaris and Trichuris are subjected to temperatures above 52°C to 54°C for even a brief period of time, most of them will lose their ability to embryonate." p. 29

* “Trichuris eggs required a more highly saturated atmosphere before they could develop than did Ascaris eggs, and the former was less resistant to desiccation. It was evident that under fractional relative humidities, the eggs of Trichuris succumbed more readily than did those of Ascaris. The explanation for the difference may lie in two basic differences in the eggs: (1) the comparative sizes: Ascaris eggs are larger and have a considerably greater surface of the fibrous membrane through which the diffusion of gases occurs; (2) the difference in time required to complete embryonation under optimum conditions: Trichuris eggs require more time to complete their development than do Ascaris." p. 30 (Nolf, 1932).

* "Many early researchers noted that Ascaris lumbricoides degenerated rapidly on various soil types when exposed to direct sunlight (Ogata, 1925; Brown, 1928; Caldwell & Caldwell, 1928; and Otto, 31 1929). Studies on the effects of humidity on Ascaris eggs under field conditions having noted that the human ascarid eggs were rapidly killed in faecal cultures on sand in the direct sun (Brown, 1928). It was concluded that desiccation and heat were both important in killing the eggs, and the results indicated that soil type is an important factor in the rate of development and viability of the Ascaris and Trichuris eggs. The cultures on sand in the sun did not produce any embryonated eggs, while those in the shade did. Those in loam, clay and humus soils became embryonated but Ascaris on humus soil was slower in development due to the minimised oxygen." p. 30

* “Studies on the effects of humidity on Ascaris eggs under field conditions having noted that the human ascarid eggs were rapidly killed in faecal cultures on sand in the direct sun (Brown, 1928). It was concluded that desiccation and heat were both important in killing the eggs, and the results indicated that soil type is an important factor in the rate of development and viability of the Ascaris and Trichuris eggs. The cultures on sand in the sun did not produce any embryonated eggs, while those in the shade did. Those in loam, clay and humus soils became embryonated but Ascaris on humus soil was slower in development due to the minimised oxygen. The temperatures on sand frequently rose above 50°C which was found to be lethal to Ascaris eggs while other soils used never reached those temperatures p. 31.” (Ogata, 1925).

* "It is quite likely that Ascaris eggs have become adapted to developing in nature in a medium, which is not fully oxygen-saturated, with the result that higher oxygen tensions are not necessary for normal development. It was originally thought that the rate of oxygen consumption was constant in embryonating Ascaris eggs (Brown, 1928); but later work by Fairbairn, (1957) showed that the rate decreases rapidly to about half its initial value during the first 36h, then increases steadily to a maximum after 10 days when the embryo is vermiform. From days 10-25 the rate decreases rapidly again, then declines more slowly to a very low level at 140 days. The initial decline was due to an oxygen debt inherited from the essentially anaerobic metabolism of the adult female worm and the second decline could be an adaptation to prolonged survival in the environment, ensuring that food reserves are not expended.”

* “The oxygen tension in faecal cultures is usually low (Fairbairn, 1957) and the initial decline could also be an adaptation to this situation. The oxygen requirements of Trichuris eggs are not essentially different from those of Ascaris eggs. Carbon dioxide given off, if allowed to remain in close contact with the eggs, will retard their development. No nitrogen is given off during development of Trichuris embryos" p. 36 (Nolf, 1932).

World Health Organization main sources:

* Feachem RG, Bradley DJ, Garelick H and Mara DD. Sanitation and Disease: Health Aspects of Excreta and Wastewater Management. John Wiley and Sons, Chichester, UK. 1983

* Nolf, L. O. (1932). Experimental studies on certain factors influencing the development and viability of the ova of the human Trichuris as compared with those of the human Ascaris. (American Journal of Hygiene, 16(1): 288-322.)

'''Further Reading and Notes:'''

* Spindler, L. A. (1929 b). [http://aje.oxfordjournals.org/content/10/2/476.full.pdf The relation of moisture to the distribution of human Trichuris and Ascaris]. American Journal of Hygiene. 10:476-496.

* Silverston NA. [https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1927001/?page=2 Whipworms in a Cambridgeshire Village]. Br Med J. 1962 Dec 29;2(5321): 1726–1728. ([https://www.ncbi.nlm.nih.gov/pmc/articles/PMC1927001/pdf/brmedj03003-0040.pdf PDF])

* Bundy DA, Cooper ES. [http://www.ncbi.nlm.nih.gov/pubmed/2683614 Trichuris and trichuriasis in humans]. Adv Parasitol. 1989;28:107-73. (Chapter 3 in: Baker JR, Muller R. [http://books.google.com/books?id=6iMDG3KURcwC&pg=PA169&lpg=PA169&dq=%22Experimental+studies+on+certain+factors+influencing+the+development+and+viability+of+the+ova+of%22&source=bl&ots=x551hJOCnv&sig=C6g7a5jFnXqJlc0htc463CPuBN4&hl=en&ei=ywqETP11woKdB5S9hWo&sa=X&oi=book_result&ct=result&resnum=6&ved=0CCgQ6AEwBQ#v=onepage&q=%22Experimental%20studies%20on%20certain%20factors%20influenc%20ing%20the%20development%20and%20viability%20of%20the%20ova%20of%22&f=false Advances in Parasitology], Volume 28 Elsevier Science. 1989.)

* Rogers WP, Summerville RI. [https://books.google.co.uk/books?hl=en&lr=&id=rS4T9Y2WTJwC&oi=fnd&pg=PA109&ots=sHlMi41pQO&sig=_5-WiUg317umcmfjVb-m5RZmv2A%23v=onepage&q&f=false The Infective Stage of Nematode Parasites and Its Significance in Parasitism Adv Parasitol]. 1963;1:109-77 (Chapter 3 in: Dawes B. [http://books.google.com/books?hl=en&lr=&id=rS4T9Y2WTJwC&oi=fnd&pg=PA109&ots=sHlMi41pQO&sig=_5-WiUg317umcmfjVb-m5RZmv2A#v=onepage&q&f=false Advances in Parsitology] vol 1.)

* Jenkins J. [http://www.scribd.com/doc/35945465/Composting-Human-Manure-Handbook The Humanure Handbook: A Guide to Composting Human Manure]. 1999.

* Feachem RG, et al. [http://www.scribd.com/doc/106288272/At-for-Water-Supply-and-Sanitation-3-Health-Aspects-of-Excreta-and-Sullage-Managment At for Water Supply and Sanitation 3, Health Aspects of Excreta and Sullage Management]. World Bank. 1980 Dec.

* Ciziulevicius, S. On the Viability of Ascaris lumbricoides and Trichuris trichiura Eggs in Compost Heaps. Acta Parasitologica Lithuanica. 1961;3:51-60. (English Summary)

* Caldwell FC & EL. [http://www.jstor.org/stable/3271383?seq=1%23page_scan_tab_contents Preliminary Report on Observations on the Development of Ova of Pig and Human Ascaris under Natural Conditions, and Studies of Factors Influencing Development]. Journal of Parasitology. 1928 Jun; 14(4):254-260

* Desiccation will kill eggs. See p. 168 in: Muller R. [https://books.google.co.uk/books/about/Worms_and_Human_Disease.html?id=bWtCMIGF_FMC&hl=en Worms and Human Disease], 2nd ed. CABI Publishing. 2002.

* Brown, H. [http://www.jstor.org/stable/3271397?seq=1%23page_scan_tab_contents Studies on the Rate of Development and Viability of the Eggs of Ascaris lumbricoides and Trichuris trichiura under Field Conditions]. The Journal of Parasitology. 1927;14(1):1-15.

* Developing eggs will die within a week, if subjected to desiccation.
# Cheng. TC. General Parasitology. 3rd ed. Academic Press. 1986, p 488.
# Garcia LS, Bruckner DA. Diagnostic medical parasitology. 2nd ed. American Society of Microbiology. 1993. p 198

* “... eggs of the genus Trichuris, which seem particularly able to survive under conditions of extreme cold.” Nakladova 1956, Hill 1957, quoted on p 141 of Dawes B. [https://books.google.com/books?id=rS4T9Y2WTJwC&pg=PA141&lpg=PA141&dq=%22Nakladova+1956,+Hill+1957%22&source=bl&ots=sJgJabVtRL&sig=UXlKvAqsXQL17-8aBd5WXnT73C4&hl=en&sa=X&ei=aCo9VdXYLMG4sAWKi4CABA&ved=0CB8Q6AEwAA Advances in Parasitology APL.] 1963.

</ref><ref follow="WhipwormEggsDevelop" group="note">
'''Whipworm eggs develop according to this schedule:'''
* 4-6 months at 15ºC (59ºF)
* 3 to 4 weeks at 26ºC (79ºF)
* 17 days at 30ºC (86ºF)
* 11 days at 35ºC (95ºF)
Muller R. [https://books.google.co.uk/books/about/Worms_and_Human_Disease.html?id=bWtCMIGF_FMC&hl=en Worms and Human Disease], 2nd ed. CABI Publishing. 2002. p 165. 
I have it in my notes that they will not develop if the temperature is over 37ºC (98.6ºF), but I don’t have the citation
for this. 

“A review of the literature furnished the following information relative to optimum temperatures for development of several species of parasitic eggs.” 
''Trichuris trichiura 30 C (86 F)'' 
Feachem et al. (1983). [http://www.bvsde.paho.org/bvsacd/cd65/SanitaryParasitology_Guide.pdf Integrated Guide to Sanitary Parasitology]. (World Health Organization, Regional Office for the Eastern Mediterranean, Regional Centre for Environmental Health Activities, Amman – Jordan, 2004. p 27. Table 2.3).

</ref><ref follow="WhipwormLiveFor" group="note">
Opinions vary widely regarding the life expectancy of whipworms in a host.
* “The adult worms normally live for about 1-2 years where there is repeated reinfection, but can survive for much longer in some circumstances.” Muller R. [https://books.google.co.uk/books/about/Worms_and_Human_Disease.html?id=bWtCMIGF_FMC&hl=en Worms and Human Disease], 2nd ed. CABI Publishing. 2002. p 165

* When reinfection does not occur, most worms die within 3 years. Myers, E. N., Negron, R. and Pearlstein, H. Intestinal parasitoses in Puerto Rican preschool children at Philadelphia, Pennsylvania in 1958. Am J Med Sci. 1959 Jan; 237(1):59–66. (cited in Marcial-Rojas, RA. 1971. Pathology of Protozoal and Helminthic Diseases. Baltimore: Williams & Wilkins. p 664.)

* Whipworms live 4-5 years. Henry JB. Clinical Diagnosis and Management By Laboratory Methods. Philadelphia: WB Saunders, 1978: 1779-88.

* Lifespan is 4 to 6 years, but can live up to 8 years. Brown, Harold W., Neva FA. 1983. Basic clinical parasitology. 5th ed. Norwalk, Conn.: Appleton-Centurry-Crofts. 339 p., p 112.

</ref><ref follow="NoCitationForThis" group="note">
I don’t have the citation for this.

</ref>
<references group="note" />

== Materials List ==

I've found that the sort of materials listed below are currently adequate for my cleaning of whipworm eggs,
although the suppliers mentioned here are not necessarily the ones I used, merely sources of similar items.
This list, as well as any items mentioned or cited previously, is offered solely for educational purposes and its
inclusion is not intended to be encouragement to anyone to emulate my practice of whipworm egg cleaning. The
Warning and Disclaimer printed above apply.

Common sense should be used when interpreting this list, because I might have been able to find substitutes for the
items listed below that would have performed the same functions. For example, when dimensions of items are given
this usually doesn’t mean I would have had to use exactly those dimensions. Such numbers are included only so the
reader has a rough idea of what I’m describing.

I have included graphics of some of the listed items for reference if and when the links expire.

=== Microscope ===
* [[Microscope selection and use]]

=== Centrifuge ===
<blockquote>
When getting a centrifuge it was important to anticipate the tubes I would need, since I had to match the tubes to
the centrifuge I bought. It turned out that I actually had to modify my tubes by cutting them off, which was a
problem, since they turned out to have harness 9, like corundum, sapphire, or ruby; so I had to use a diamond tile
saw. I purchased a used centrifuge locally, but, if I hadn’t been able to, I could have bought one on eBay. The
centrifuge I have is a horizontal one, which is supposed to result in a cleaner separation line, but I’m not sure how
much difference this makes. Its relative centrifugal force ([https://en.wikipedia.org/wiki/Centrifuge#Mathematical_description RCF]) is about 1,600 times gravity, and its speed is about
4,000 RPM. So, if I were to use only Earth's natural gravity, it would take 10 minutes times 1,600 to perform this
separation. So 10 minutes X 1,600 = 16,000 minutes, which works out to about 11 days.

I found these guides useful:
* [http://www.ebay.com/gds/How-to-Buy-Centrifuges-/10000000177627198/g.html How to Buy Centrifuges by eBay]
* [http://www.druckerdiagnostics.com/centrifuge/centrifuge-selector.html Centrifuge Sector Guide]
* [http://www.coleparmer.com/TechLibraryArticle/30 Basics of Centrifugation]
* [http://www.woodleyequipment.com/laboratory-diagnostics/clinispin-centrifuges-171.html Why Horizontal Separation?]

Budget centrifuges:
* Scientists at Harvard have developed a manually-operated, light and portable, very inexpensive centrifuge.
** [https://gmwgroup.harvard.edu/pubs/pdf/1033.pdf Egg beater as centrifuge: isolating human blood plasma from whole blood in resource-poor settings]
* [http://www.sciencedaily.com/releases/2010/05/100503135713.htm Salad spinner useful to separate blood without electricity in developing countries]

</blockquote>

=== Lab Equipment ===

<blockquote>
<div style="margin:12px;">
[[file:TTO-Inc002.jpg|link=]]
</div>
{{Columns|3}}
* '''Centrifuge Tubes''' e.g., [http://www.camlab.co.uk/centrifuge-tubes-c2792.aspx Centrifuge Tubes, plastic] (see graphic 2A)

* '''Test Tube Rack''' I had to be careful to match the size of my test tube rack to my test tubes. I really like the type shown here (see graphic 2B). e.g.
** [http://www.globescientific.com/rack-tube-16-17mm-60-place-pp-white-p-1143.html?sku_id=456500 Globe Scientific Inc., 16/17 mm, 60 tube capacity]
** [http://www.seohcorp.com/seoh-test-tube-rack-plastic-for-60-tubes-16mm/ SEOH TEST TUBE RACK PLASTIC For 60 tubes 16mm] (see graphic 2B)

* '''Test tube brush, small''' e..g., [http://www.hometrainingtools.com/test-tube-brush Test tube brush, small] (see graphic 2C)

* '''Minipipette, 100 microlitre''' e.g.
** [http://www.rakuten.com/prod/100-microliter-li-lpet-mini-pipette-pipettor/293178581.html Mini Pipette Pipettor]
** [http://www.onlinesciencemall.com/100-microliter-li-lpet-mini-pipette-pipettor.html 100 Microliter "Li'lpet" Mini Pipette Pipettor] (see graphic 2D)
** [http://www.carolina.com/biotechnology-pipets/fixed-volume-100-ul-03-ulminipipet/214304.pr?question=214304 Fixed-Volume 100 µL (±0.3 µL) MiniPipet] (see graphic 2E)

* '''Minipipette tips''' When I bought my mini pipette tips I had to check carefully to make sure that they were compatible with my particular mini pipette, e.g.
** [http://www.membrane-solutions.com/Pipette_micro_tips.htm MS® Pipette Tips]
** [http://www.gilson.com/en/Pipette/Categories/55/%23.VcjANnhcuFU Diamond Tips - Gilson]
** [http://www.labx.com/v2/lablinks/linkscategory.cfm?catID=122 Pipette Tips Company Links]
** [http://www.thomasnet.com/products/pipettes-58671009-1.html Pipettes Suppliers]
** [http://www.biohit.com/resource/files/media/brochures/liquid-handling/all/Tips_insert_optifit2009_SCREEN.pdf Biohit Optifit Tips] (see graphic 2F)
** [http://www.onlinesciencemall.com/pk-100-1-200-microliter-universal-fit-micropipette-tips.html Pk/100 1-200 Microliter Universal Fit Micropipette Tips, Natural Color] (see graphic 2G)

* '''Pasteur pipettes''' (see graphic 2H) e.g.
** [http://www.arssales.com/epe-disppipette.html Disposable 1.0 ml Transfer Pipettes]
** [http://www.hometrainingtools.com/pipet-disposable-1-ml-10-pk/p/CE-PIPET/ Pipette, disposable, 5ml, 10 pk]
** [http://www.camdengrey.com/essential-oils/pipettes-8-3-quarters-inches-bulk.html Pipette, disposable, 8-3/4 in. long]

* '''Graduated cylinder''' e.g., [http://www.hometrainingtools.com/graduated-cylinder-poly-250-ml/p/CE-CYPP250/ Graduated cylinder, poly, 250 ml and 500 ml: item# CE-CYPP250] (see graphic 2I)

* '''Hydrometer''' e.g.
** [https://www.wardsci.com/store/catalog/product.jsp?catalog_number=150884 Wards Natural Science Specific Gravity Hydrometer 1.00 - 2.00 model] (see graphic 2J)

* '''pH Meter, Digital, 0-14 pH range''' e.g. [http://www.hometrainingtools.com/ph-meter-digital-0-14-ph-range Milwaukee pH600 pocket digital pH meter / 0-14 pH range] (see graphic 2K)

* '''pH 7.0 Calibration Solution''' If I happen to have run out of calibration fluid I will calibrate my pH meter by using distilled water, which should be about 7 pH, and newly bought pH 3 battery acid.
** e.g. [http://www.hometrainingtools.com/ph-7-0-calibration-solution-5-packets pH 7.0 Calibration Solution, 5 packets] (see graphic 2L)

* '''Small capacity postal scale''' e.g.
** [http://www.amazon.co.uk/Helix-Salter-FC3250-500x10g-Foldable/dp/B00KG82WUC/ref=sr_1_28?ie=UTF8&qid=1437761247&sr=8-28&keywords=postal+scale+mechanical Helix Salter FC3250 500x10g Foldable Postal Letter Scale]
** [http://www.amazon.co.uk/gp/product/B00FK3VRCC?keywords=postal%20scale%20mechanical%202%20lb&qid=1437760984&ref_=sr_1_2&sr=8-2 Smart Weigh TZ5000 Sleek Cuisine Stainless Steel Digital Kitchen Scale 5000g X 1g] (see graphic 2M).
{{Columns}}

<div style="margin:12px;">
[[file:TTO-Inc003.jpg|link=]]
</div>

{{Columns|3}}
* '''Small funnel''' I don't currently need to use this size of funnel, but this is the size I would need if, for some reason, I needed to fill test tubes. This type of funnel is typically used for filling perfume bottles, and its neck is small enough to fit into the tube's mouth. e.g., [http://www.onlinesciencemall.com/plastic-funnel-65mm-polypropylene-long-stem.html Plastic Funnel: 65mm Polypropylene Long Stem Pk 1] (see graphic 3A).

* '''Larger Funnel''', for filling the Chug bottles e.g., [http://www.amazon.co.uk/LARGE-FOOD-GRADE-FUNNEL-HOMEBREW/dp/B005YZP1P0/ref=sr_1_20?s=kitchen&ie=UTF8&qid=1437528290&sr=1-20&keywords=Funnel+kitchen 1 x 9" LARGE FOOD GRADE FUNNEL] (see graphic 3B)

* '''Set of three nested funnels.''' The largest of these is probably the right size for filling sprayer bottles. e.g., [http://www.amazon.co.uk/gp/product/B00175HEXS?keywords=kitchen%20funnel&qid=1437528886&ref_=sr_1_1&sr=8-1 Chef Aid 3-Piece Funnel Set, White] (see graphic 3C).

* '''Sieve''' e.g., [http://www.amazon.co.uk/Premier-Housewares-Sieve-Stainless-Steel/dp/B0052R78JC/ref=sr_1_1?s=kitchen&ie=UTF8&qid=1437338495&sr=1-1&keywords=sieve+colander Premier Housewares Sieve - 20 cm - Stainless Steel] (see graphic 3D). This model looks a lot like the one I bought, before I cut the handle and prongs off with my hacksaw.

* '''Meshes'''
# [http://www.amazon.com/Nylon-Opaque-Off-White-Length-microns/dp/B001D8GD4U/ref=sr_1_1?s=industrial&ie=UTF8&qid=1428015733&sr=1-1&keywords=30+micron+mesh Nylon 6/6 Woven Mesh Sheet, Opaque Off-White, 12" Width, 12" Length, 30 microns Mesh Size, 18% Open Area (Pack of 5)] (see graphic 3E).
# [http://www.amazon.com/Nylon-Opaque-Off-White-Length-microns/dp/B001D8ECDE/ref=sr_1_2?s=industrial&ie=UTF8&qid=1428015971&sr=1-2&keywords=20+micron+mesh++5 Nylon 6/6 Woven Mesh Sheet, Opaque Off-White, 12" Width, 12" Length, 20 microns Mesh Size, 14% Open Area (Pack of 5)]

* '''Clips for holding the meshes''' e.g.
** [http://www.amazon.co.uk/Kitchen-Craft-Magnetic-Memo-Clips/dp/B000YJDK68/ref=pd_bxgy_201_img_z Kitchen Craft Set of 4 Magnetic Memo Clips] (see graphic 3F). Although the magnets on these clips are rather weak, they’re just strong enough for my purposes.
** [http://www.amazon.co.uk/Kitchen-Craft-Magnetic-Clips-Multi-Colour/dp/B00U1A0LHG/ref=sr_1_4?ie=UTF8&qid=1438106755&sr=8-4&keywords=magnetic+bag+clips Kitchen Craft Magnetic Bag Clips, Multi-Colour]
** [http://www.amazon.co.uk/OXO-Good-Grips-Magnetic-Multi-Colour/dp/B0007P4J3M/ref=sr_1_5?ie=UTF8&qid=1438107081&sr=8-5&keywords=Magnetic+Bag+Clips OXO Good Grips All Purpose Magnetic Clips, Multi-Colour]

* '''Trash container''' (lid type) e.g., [http://www.amazon.co.uk/Brabantia-Pedal-Bin-Plastic-Bucket/dp/B00CDKH2RO/ref=sr_1_1?s=kitchen&ie=UTF8&qid=1437527903&sr=1-1&keywords=plastic+Trash+container+%28lid+type%29 Brabantia Pedal Bin with Plastic Bucket, 3L - Brilliant Steel] (see graphic 3G)

* '''Stock pot''' 11 litres (12 quarts) e..g., [http://www.amazon.co.uk/Buckingham-Stock-Pot-Stainless-Steel/dp/B0049MPIM0/ref=sr_1_3?s=kitchen&ie=UTF8&qid=1437527961&sr=1-3&keywords=Stock+Pot Buckingham Stock Pot with Stainless Steel Lid 26 cm, 11 L] (see graphic 3H).
{{Columns}}

<div style="margin:12px;">
[[file:TTO-Inc004.jpg|link=]]
</div>

{{Columns|3}}
* '''Plastic containers''' - 0.5 litre (16 fl oz) - for holding bleach or ammonia, and for use when neutralising the car battery acid.

* '''Plastic containers''' - 0.5 litre (16 fl oz). A small supply of approximately pint-sized containers (to hold the cultures) with holes in their bases for drainage. Plastic tomato plant starters, or similar, would have been suitable, but I made my own by drilling holes in the bottoms of plastic food containers. I am very likely being too pernickety but, for all the equipment that I use to hold liquids or that will come into contact with moisture, I avoid using plastics that might contain Bisphenol A (see [http://www.healthiertalk.com/why-all-fuss-about-bisphenol-bpa-1043 Why All the Fuss About Bisphenol A (BPA)]) so I stick with plastic types 1, 2, 4, and 5. I also tried to keep to these types of plastic for other items that I used for this protocol - at least when I could tell what the plastic type was.

* '''Plastic containers''' - assorted, to hold miscellaneous items e.g. 2 litre (64 oz), 15 cm x 20 cm x 9 cm (6” x 8” x 3½”) and 2 x 15 litre (16 quart) containers
** e.g., [http://www.amazon.co.uk/gp/product/B00CBX3XQW?keywords=15%20litre%20food%20storage%20rubbermaid&qid=1437594425&ref_=sr_1_fkmr1_3&sr=8-3-fkmr1 15 Litre Plastic Storage Box Container With Clip On Lid and Handle] (see graphic 4A).

* '''Soft, thin plastic bowl''' that has the top 75% cut off, leaving the bottom, and thereby making my hand scoop. e.g., [http://www.amazon.co.uk/Colourworks-Melamine-Bowls-Set-Four/dp/B003WOJ6FQ/ref=sr_1_3?s=kitchen&ie=UTF8&qid=1437528109&sr=1-3&keywords=Bowl+plastic+cereal Colourworks Melamine Bowls, 15 cm (Set of Four)] (see graphic 4B)

* '''Glad containers''' e.g.
** [https://www.glad.com/food-storage/containers/mini-round/ Mini Round] (see graphic 4C)
** [https://www.amazon.co.uk/d/Home-Kitchen/Glad-Mini-Round-Containers/B000F2NJBA Gladware Mini Round 4 oz Containers with Lids]

* '''Yogurt container/cup''', e.g. (see graphic 4D)

* '''Party cups''' e.g., [http://www.amazon.co.uk/Ruby-Apple-American-Party-Cups/dp/B00LI2MGZ6/ref=sr_1_1?s=kitchen&ie=UTF8&qid=1437528258&sr=1-1&keywords=Cups Ruby Apple Red American Party Cups - 16oz (455ml) - Disposable Party Cups - Packs of 50 or 100 (50)] (see graphic 4E).

* '''Shot glasses''', 2 – 4, either plastic or glass e.g., [http://www.amazon.co.uk/Plastic-Shot-Glasses-30ml-total/dp/B001IZVS2A/ref=pd_cp_k_h_b_cs_0 Plastic Shot Glasses 30ml 30/Pack] (see graphic 4F), or [http://www.shot-glasses.co.uk/categories.php?cat=95 Shot-glasses.co.uk]

* '''Chug Style Bottle''' 32 ounce/1 quart Rubbermaid "Chug bottles". I need at least two of these. e.g.
** [http://www.amazon.co.uk/Rubbermaid-SippN-Sport-Style-Bottle/dp/B000RNHODQ/ref=sr_1_1?ie=UTF8&qid=1419291373&sr=8-1&keywords=Chug+bottles Rubbermaid 1 Qt. Sipp'N Sport Chug Style Bottle]
** [http://www.wayfair.com/Rubbermaid-1-Quart-Chuggables-Bottle-3163RDEDAY1-QU2591.html Wayfair.com 1 Quart Chuggables Bottle] (see graphic 4G)

* '''Spray bottle''' (for dechlorinated water). I use 6 of these e.g.
** [http://www.rubbermaid.com/en-US/heavy-duty-spray-bottle Rubbermaid Heavy-Duty Spray Bottle] (see graphic 4H)
** [http://www.amazon.co.uk/Green-Blade-Liquid-Bottle-Gardening/dp/B000QF7IB8/ref=sr_1_3?ie=UTF8&qid=1318522047&sr=8-3 Green Blade 750ml Water & Liquid Spray Bottle].

* '''Used plastic pill bottles''', miscellaneous, for sharps, etc.

* '''Bin organisers''' - for small items. Any of these would work well:
** [http://www.amazon.com/Stack-CB-12-Clear-12-Bin-Organizer/dp/B000M2WVT4 Stack-On CB-12 Clear View 12-Bin Organizer]
** [http://www.amazon.com/Gladiator-GarageWorks-GAWESB6PSM-Small-6-Pack/dp/B000Q38P7Q/ref=sr_1_1?s=hi&ie=UTF8&qid=1314898562&sr=1-1 Gladiator GarageWorks GAWESB6PSM Small Item Bins, 6-Pack]
** [http://www.spectrum-scientifics.com/SearchResults.asp?Search=Lab+Supply+Bin+for+Small+Items+-+13+Compartments+%231922 Lab Supply Bin for Small Items - 13 Compartments]
** Mine look something like these: [http://www.amazon.co.uk/Faithfull-Plastic-Storage-Mounting-Pieces/dp/B004UR2GFQ/ref=sr_1_25?ie=UTF8&qid=1318522302&sr=8-25 Faithfull Plastic Storage Bins with Wall Mounting Rails (12 Pieces)] (see graphic 4I)

* '''Mini fridge''' (see graphic 4J) I store the filled Glad containers in an unplugged mini fridge e.g., the Caldura 17 litre Compact Mini Fridge (see graphic 4I) on the assumption that, if my home were to burn down, my precious stock of eggs would have a better chance of survival if protected by the fridge's insulation. I also keep my important papers, including inoculation records, in the fridge. As I typically open this fridge about once each day to retrieve some paper or other, this arrangement ensures that the air in the fridge circulates periodically, although the eggs must use very little air.

* '''Plastic washing up bowls''' I have used three of these in the past (approximately 10 cm deep x 30 cm x 30 cm [4” deep x 12” x 12”]) and they can still come in handy, but I now generally use the sink for all washing purposes. For example, when I rinse the meshes I run these under the flowing water from the tap. Then I either hang them up, or, if I'm going to use them right away, spray rinse them with the dechlorinated water. e.g.
** [http://www.hardwareandtools.com/rubbermaid-home-2951-ar-wht-white-rectangle-dishpan-egba-3249.html Rubbermaid Home 2951-Ar Wht White Rectangle Dishpan]
** [http://www.amazon.co.uk/Casa-Rectangular-Washing-Up-Bowl/dp/B002SLCPH0/ref=sr_1_6?s=kitchen&ie=UTF8&qid=1318522411&sr=1-6 Rectangular Washing Up Bowl – Cream]
** [http://www.amazon.com/Medline-Plastic-Graduated-Rectangular-Unboxed/dp/B000WAW966 Medline Plastic Graduated Rectangular Wash Basin, Gold, Unboxed(single Unit) by Medline]
** [http://www.amazon.co.uk/Casa-Rectangular-Washing-Up-Bowl/dp/B002SLCPH0/ref=sr_1_6?s=kitchen&ie=UTF8&qid=1318522411&sr=1-6 Rectangular Washing Up Bowl – Cream by Whatmore] (see graphic 4K)

* '''Dishrack''' - I find it useful to have a separate one for drying washed lab equipment.

* '''Bluettes''' - [https://www.amazon.co.uk/Lehigh-Spontex-17005-Bluettes-Rubber-x/dp/B000UGRO7G Lehigh Spontex 17005 Bluettes Knit Rubber Glove] (see graphic 4L).

* '''Disposable exam gloves in latex, nitrile or vinyl''' It’s far cheaper to reuse the gloves a number of times, but if these are put back on too soon, they may still be damp from the previous use and hard to get back on. In this case, I use a hair dryer to dry them out. Alternatively, I may place the dryer on my wrist, while it’s set on cool rather than hot, and use it to inflate the glove (which at that moment is stretched over both my wrist and the dryer’s mouth) as I work the glove back on. Yet another option is to rotate several pairs. These are widely available at local pharmacies and online, e.g.
** [http://www.amazon.co.uk/Simply-Powder-Free-Latex-Gloves-Medium/dp/B004CFR5VQ/ref=sr_1_7?ie=UTF8&qid=1319328987&sr=8-7 Simply Powder-Free Latex Gloves Medium - 100-pack] (see graphic 4L)
** [http://www.amazon.co.uk/Powder-Gloves-Disposable-Medical-Medium/dp/B00M13JGUO/ref=sr_1_2?ie=UTF8&qid=1430596949&sr=8-2&keywords=disposable+exam+gloves 200 (2 Box) x Vinyl Powder Free Gloves Disposable Clear Food Medical etc. (Medium)]
** [http://www.amazon.co.uk/Bodyguards-Clear-Powder-Gloves-Medium/dp/B0086V8124/ref=sr_1_1?ie=UTF8&qid=1430597934&sr=8-1&keywords=latex-free+exam+gloves Bodyguards Clear Vinyl Powder Free Exam Gloves Medium Box of 100]

* '''Vinyl lab apron''' I found mine at an online science supply site. Some college bookstores may also carry these for their chemistry students. On mine, the apron’s tie cords were made of plastic, and quickly broke, so I replaced them with used shoe laces attached to the apron with shoe goo and staples. e.g.
** [http://www.ebay.co.uk/itm/Black-apron-water-proof-resist-vinyl-back-chef-cook-butchers-pocket-bib-halter-/231354595136?pt=LH_DefaultDomain_3&var=&hash=item35ddcef740 Black apron water proof resist vinyl back chef cook butchers pocket bib halter]
** [http://www.hometrainingtools.com/lab-apron-no-2-heavy-vinyl/p/CE-APRON2/ Lab Apron No. 2, heavy vinyl
** [http://www.hamiltonbell.com/product-catalog/item/lab-aprons-27x42 Hamilton Bell Co. Inc., No. 5250, 27” x 42”] (see graphic 4N).

* '''Shoe goo''' - [http://www.amazon.co.uk/Original-Shoe-Goo-CLEAR-110ml/dp/B000BQR55G/ref=sr_1_1?s=kitchen&ie=UTF8&qid=1437527367&sr=1-1&keywords=Shoe+Goo Original Shoe Goo CLEAR - 110ml/3.7oz Tube]

* '''Scissors with a fine point''' e.g.
** [http://www.amazon.co.uk/dp/B00YMWW3XE/ref=sr_1_13?ie=UTF8&qid=1436919223&sr=8-13&keywords=Scissors+Fine+Point Nurses Scissors Sharp/Sharp points (331-00)]
** [http://www.amazon.co.uk/MagiDeal-Stainless-Pointed-Eyelash-Scissors/dp/B01147AARK/ref=sr_1_51?s=beauty&ie=UTF8&qid=1436921747&sr=1-51&keywords=pointed+scissors MagiDeal Stainless Steel Pointed Tip Eyelash Trimmer Eyebrow Scissors] (see graphic 4P)

* '''Hacksaw''' e.g., [http://www.amazon.co.uk/Stanley-Dynagrip-Heavy-Duty-Hacksaw/dp/B0001GRVB4/ref=sr_1_4?ie=UTF8&qid=1437504905&sr=8-4&keywords=hack+saw Stanley Dynagrip Heavy Duty Hacksaw 1 20 110] (see graphic 4Q).

* '''Kitchen sponges''', several e.g. [http://www.amazon.co.uk/gp/product/B00006IBMC?keywords=O-Cel-O%20Sponges,%204%20Count&qid=1437594134&ref_=sr_1_1&sr=8-1 3M O-Cel-O Handy Sponge Power Pack 7274T, 4-Count] (see graphic 4R). I really like this type of sponge for making a bed to rest my hydrometer on.

* '''Rubber bands''' e.g. [https://www.amazon.co.uk/d/Office-Supplies/UNIVERSAL-OFFICE-PRODUCTS-Rubber-Universal%C3%83%C6%92%C3%82%C3%83%C3%82-/B018ORH4CM Universal Rubber Bands, Size 16, 1lb Pack by Universal] are about right

* '''Zip-lock Bags''' - [https://www.amazon.co.uk/gp/product/B004DITJRK Ziploc 17.7cm x 19.5cm bags] (see graphic 4S)

* '''Disposable plastic spoons''' e.g., [http://www.amazon.co.uk/Disposable-Plastic-Quality-Wedding-Cutlery/dp/B0066R1DS4/ref=sr_1_1?s=kitchen&ie=UTF8&qid=1437528282&sr=1-1&keywords=Plastic+spoons 100 x Disposable White Plastic High Quality Spoons For Party / Wedding Cutlery] (see graphic 4T)
{{Columns}}

<div style="margin:12px;">
[[file:TTO-Inc005.jpg|link=]]
</div>
{{Columns|3}}
* '''Rid-x''' - [https://www.amazon.co.uk/Rid-X-Septic-System-Treatment-Supply/dp/B0058I168Y/ref=lp_5326162031_1_1_a_it RID-X Septic Tank System Treatment, 3 Month Supply Liquid, by Rid-X] (see graphic 5A) 

* '''Bleach''', by the litre (or gallon), 5-6% sodium hypochlorite (Or) '''ammonia''', by the litre (or gallon), 2-3%

* '''Car battery acid''' (sulphuric/sulfuric) e.g., [http://www.amazon.co.uk/gp/product/B00NMKSZGG?keywords=car%20battery%20acid&qid=1437341267&ref_=sr_1_10&sr=8-10 Battery Acid 1 Litre Electrolyte for Dry Batteries car motorbike SULPHURIC] (see graphic 5B)

* '''Distilled water''' e.g.
** [http://www.amazon.co.uk/gp/product/B0035FF28M?*Version*=1&*entries*=0 Distilled Water - 5.5 Litres]
** [http://www.buydistilledwater.co.uk/distilled-water/3-5-litre-distilled-water.html 5 Litre Distilled Water]
** [http://www.amazon.co.uk/The-Water-Delivery-Company-Distilled/dp/B003Q98QMG/ref=sr_1_2?s=kitchen&ie=UTF8&qid=1437528145&sr=1-2&keywords=Distilled+water Distilled Water - 20 litres] (see graphic 5C)

* '''Terry cloth towels''', 2 or 3, small
* '''Paper towels''', 1 roll
* '''Plastic refuse bags''' for triple bagging and disposing of old soil and stool

* '''Sugar (sucrose)''', a number of bags e.g., [http://www.amazon.co.uk/Tate-Lyle-Granulated-Sugar-1KG/dp/B00AYY5DBS/ref=sr_1_7?s=grocery&ie=UTF8&qid=1438107762&sr=1-7&keywords=sugar Tate & Lyle Granulated Sugar 1KG] (see graphic 5D).

* '''Baking soda''' (sodium bicarbonate/sodium hydrogen carbonate/[https://en.wikipedia.org/wiki/Bicarbonate NaHCO3])
** e.g., [https://www.amazon.co.uk/Arm-Hammer-Pure-Baking-Soda/dp/B01BFGV4E8 Arm & Hammer Baking Soda 454 g] (see graphic 5D)
{{Columns}}
</blockquote>

== Helminth incubation discussion groups ==

* [[Helminth incubation#Helminth incubation groups | Helminth incubation groups]]

== Suggestions/observations ==

If anyone using the information on this page has any suggestions for its improvement, or any other observations about it, please post these to any of the following groups.
* [https://www.facebook.com/groups/678894952216125/ Facebook Helminth Incubation group]
* [https://www.facebook.com/groups/htsupport/ Facebook Helminthic Therapy Support Group]
* [https://groups.yahoo.com/neo/groups/helminthictherapy/info Yahoo Helminthic Therapy Forum]
They will then be collected and added to this page.

User talk:Alana

2017-10-13T22:08:57Z

Alana: /* Chefast Hydrometer */ new section

Hi Alana, another wiki User (Tracy) wants to put a couple of questions to you about your NA and TTO protocols, so I’ve suggested that she uses the wiki to do this because it allows wiki Users to communicate with one another via the email addresses they signed up with, without actually revealing those addresses to anyone. Only the wiki software knows what these are.

I’ve sent a message to Tracy, telling her what to do to contact you this way, and you should be able to read my message to her here: http://helminthictherapywiki.org/wiki/index.php/User_talk:Tracy You can also write on her Talk page, if you want to, after hitting the “Edit” button at the top right of the page.

Once you've finished your message, add four tildes (~), which the wiki will replace with your name, as a signature, after you’ve hit the “Save page” button.

If Tracy sends you a message using your Talk page, you can reply to her on this page, after hitting the "Edit" button.

If you want to remove any messages, once they've been seen, just re-enter the page using the "Edit" button, highlight/delete the comment, and hit "Save page". [[User:John|John]] ([[User talk:John|talk]]) 03:48, 18 September 2017 (UTC)

== Clarification of Your Incubation method for TTO and NA ==

Hello.

I have read through your TTO protocol (thank you very much for creating such detailed document for the wiki) and just need a bit of clarification about your incubation protocol.

Question 1 - I understand that many use Sarah's method for incubating NA but would like to know what your practice is, please.

Question 2 -have you ever found a research paper about incubating TTO? I suspect not as you have included it in your thorough cited sources. I just wanted to double check.

I am going to attempt to incubate and process my own TTO and was hoping I could get this bit of clarification.

My background. I have been using TSO and NA and TTO to keep Crohn's Disease in remission for the past 11 years and have experimented with them to see what works best and TTO and TSO work best for me. TSO is prohibitive because of its cost and fortunately, TTO has worked very well for me. I also use NA for its systemic effect and I think it works well as an added buffer against autoimmune response activation, in general.

John Scott suggested I contact you. So here I am. Thank you in advance for your suggestions.

[[User:Tracy|Tracy]] ([[User talk:Tracy|talk]]) 19:17, 2 October 2017 (UTC)

==Testing==

I'm checking to see whether you get a notification of this message, Alana.

[[User:John|John]] ([[User talk:John|talk]]) 02:30, 3 October 2017 (UTC)

Hi John. I'm responding to your message. I received an email notification. [[User:Tracy|Tracy]] ([[User talk:Tracy|talk]]) 16:33, 3 October 2017 (UTC)

== Hello John and Tracy ==

This is just confirming that I'm now able to use this forum.
[[User:Alana|Alana]] ([[User talk:Alana|talk]]) 21:20, 13 October 2017 (UTC)

Hi Alana & John! This is wonderful.

Alana, Thank you for your input so far. Your protocol is wonderfully detailed. I am currently making a list of all the materials I will need and referencing it with the materials at the lab.
Fortunately, the bio-hackers that formed and created the lab space are keen adventurous university grad students and researchers at a local hospital, making access to reagents and other medical grade supplies possible.

It should take couple more weeks to get underway, I need to order a few things - 20 micron nylon mesh is expensive!

Hydrometer question: Can I use a hydrometer for brewing? I would like to if I can because I brew kombucha. https://www.amazon.com/dp/B0735B5YND?psc=1

[[User:Tracy|Tracy]] ([[User talk:Tracy|talk]]) 21:49, 13 October 2017 (UTC)

== Chefast Hydrometer ==

Since I haven't used this brand of hydrometer before I needed to take a quick look at the protocol and be sure that this hydrometer covered the full range of densities discussed in the paper. If it did then I don't see that there should be any problems. So https://www.amazon.com/dp/B0735B5YND?psc=1 says, "Measuring specific gravity from 0.99-1.17" Sorry, but you need to be able to go up to 1.270 for the Sheather's solution. In the paper I list "Wards Natural Science Specific Gravity Hydrometer 1.00 - 2.00 model." (I also just now realized I mistakenly included one that wouldn't have worked. "150879 Specific Gravity Hydrometer, 1.00–1.20, 0.002 divisions (see graphic 2J)") [[User:Alana|Alana]] ([[User talk:Alana|talk]]) 22:08, 13 October 2017 (UTC)

User talk:Alana

2017-10-13T21:20:41Z

Alana: /* Hello John and Tracy */ new section