The Brewing-Science Institute's
Brewers' Laboratory Handbook:
BREWING WITHOUT THE BLINDFOLD™

711 North Hancock Avenue |Colorado Springs CO 80903 |(719) 460-0418 (FAX) 475-8206 |www.brewingscience.com

"Brewing without the blindfold" means keeping an eye on your microbes, be they brewing yeast or the myriad contaminants that plague breweries. This handbook helps you do just that, and respects the fact that you want to spend time in the brewery, not in the lab!

Overview of Microbiological Methods Used in the Brewery

Aseptic Technique

The ability to transfer cultures without contamination is imperative to good microbiology. Keep in mind:

1. Hands must be washed and work areas must be cleaned with a 5% solution of disinfectant
2. Sterile vessels and liquids must be used (subject to 15psi for 15 minutes in a pressure cooker)
3. Sterile transfer tools must be used (dip in 70% ethanol or rubbing alcohol and flame)
4. Aerial fall-out and drafts must be minimized.

See Protocol: Drawing Samples, Protocol: Swabbing and Protocol: Plating Samples.

Quick & Dirty "Film Test" for Bottled Beer

Bottled products can be checked for gross contamination without much difficulty. The following are undesirable:

1. Bottle necks with a rings at the beer/headspace interface
2. Cloudiness in any part of the body
3. Residue at the bottom of filtered beer
4. Excessive foaming of opened, chilled beer.

Odor and taste are indicators as well. Most useful is catching contamination well before it gets to taste, odor and visibility thresholds.

Microscopic Examination

For yeast, use the 40x lens and make note of:

1. percent viability
2. uniformity of size and shape, or lack thereof
3. excessive amounts of trub in the suspension.

For bacteria, use the oil immersion lens and make note of:

1. whether cells are rods (bacilli) or rounds (cocci)
2. the approximate length or diameter in microns
3. whether the cells form pairs, chains or clusters
4. the gram reaction

Microscopic examination of samples has limited use. It is not helpful for detecting even moderate levels of contamination, whether it be in yeast, wort or beer. If enough bacteria or wild yeast is present to be seen while surveying only a few fields, contamination levels are very high. Also, different yeast strains can have very similar appearances under the microscope, so it can be impossible to tell a wild strain from a brewing strain, or to tell one brewing strain from another.

Detecting Mixed Strains of Brewing Yeast

If you use more than one strain of yeast in the brewery, you may wish to determine whether or not they are becoming intermingled. By plating the slurry for single colonies on nutrient medium and incubating for 5-7 days, you will be able to tell one from another after careful examination of colony shapes, sizes and colors. Any differences in these characteristics are solid indications of different strains.

See Protocol: Pouring Plates and Protocol:Streak-Plating for Single Colonies / Strain Purification

Detecting Wild Yeast, Petite Mutants & Bacteria with Selective Media

If you know what type of contaminant you’re encountering, you are better equipped to determine how serious a problem it presents and how to get rid of it. On a non-selective medium, such as nutrient agar, yeast colonies are relatively large, opaque, whitish and waxy-looking, while bacteria colonies are generally smaller, more translucent and slick-looking. Knowing this is fine, but a selective medium is needed for more specific identification. We recommend the following selective media for the basic groups of brewing contaminants:

1. wild yeast – LCSM and LWYM
2. petite mutants – nutrient agar containing or over-layed with TTC
3. brewing bacteria – SDA.

Each of these media are easy-to-use and allow for enumeration (counting the number of contaminants in the sample tested) and differentiation (knowing one type of organism on the plate from another), which is more than can be said for most testing media. Suppliers and prices are listed in the lab products section of the BSI website.

See Protocol: Pouring Plates, Protocol: Plating Samples, and Protocol: Serial Dilutions / Streak-Plating for Single Colonies / Strain Purification.

Simple Tests for Identifying Bacteria

Make the first 4 observations of any bacteria colonies before disturbing the colony with the gram stain or other tests:

1. aerobic and/or anaerobic growth (if incubated other than aerobically)
2. odor (vinegary, rotten, sulfury, fruity)
3. acid production (halo surrounding colony, with color change and/or clearing of medium)
4. colony color, size, texture and shape
5. gram reaction
6. catalase and oxidase reactions.

The gram stain takes advantage of certain differences in cell membrane properties of bacteria. All bacteria are divided into two categories: gram-positive (takes on crystal violet stain and turns blue) and gram-negative (takes on safranin stain and turns pink). Yeasts are gram-positive. The catalase and oxidase reactions test for the presence of certain enzymes which are useful for differentiating among bacteria. Armed with all the above information, check Table of Brewing Bacteria for bacteria’s identity.

See Protocol: Gram Staining and Protocol: Catalase & OxidaseTests.

Cell Counts and Determination of Proper Pitching Rate

A hemacytometer is essential for accurate cell counts (number of yeast cells/ml in a suspension). The average number of cells on a chosen grid is converted into cells/ml. This number is compared to the proper pitching rate.

See Protocol: Cell Counting.

Protocols

Protocol: Where & When to Sample

SAMPLE	FREQUENCY	SAMPLE SIZE	COMMON CONTAMINANTS	TOLERANCE*
water supply	1/week	100ml filtered	enteric bacteria, molds	<10 cfu
wort	every brew	1.0ml	enteric, acetic & lactic bacteria, wild yeast	<10 cfu; 0 cfu wild yeast
pitching yeast	every crop	1.0ml	enteric, acetic & lactic bacteria, wild yeast	<10 cfu; 0 cfu wild yeast
fermenting beer, days 1-2	every tank	1.0ml	enteric, acetic & lactic bacteria	<10 cfu; 0 cfu wild yeast
fermenting beer, days 3-5	every tank	1.0ml	acetic & lactic bacteria, wild yeast	<10 cfu; 0 cfu wild yeast
storage tank	3/week	1.0ml	acetic & lactic bacteria	<10 cfu
finishing tank	3/week	1.0ml	lactic bacteria	<10 cfu
bottling tank	1/month	100ml filtered	lactic bacteria	<10 cfu
bottled beer	every batch	100ml filtered	acetic & lactic bacteria	<10 cfu
CIP’d surfaces	every CIP	swab	-	0 cfu
*cfu = colony-forming units, or the number of colonies growing on the test plate

Protocol: Drawing Samples

Equipment

• 70% ethanol or rubbing alcohol
• sterile sample tube

Procedure

• Wash and dry hands thoroughly.
• Sterilize intervening "outside" surfaces, such as valves, with 70% alcohol and flame.
• Do not uncap tube until immediately before sampling.
• Do not touch inside of tube or cap to any surface.
• If possible, allow sample to flow or crumble directly into tube without using utensils.
• Do not allow sample to overflow onto outside of tube.
• Recap tube as soon as possible.

Protocol: Swabbing

Equipment

• sterile swab tube

Procedure

• Wash and dry hands thoroughly.
• Unscrew swab/cap from tube without touching inside of tube or cap to any surface.
• Firmly swab area to be tested, twisting swab to expose entire tip to area in question.
• Immediately place exposed swab back into tube, tighten cap and label.

Notes:

• Exposed swabs can be tested by
  
  streaking across the surface of plated media (see Protocol: Plating Samples
  or by adding 10ml sterile wort to swab tube and allowing to grow in a warm area (~86°F/30°C) for 3 days (see Protocol: Wort Stability Test).

Protocol: Wort Stability Test

Equipment

• 70% ethanol or rubbing alcohol
• sterile sample tube

Procedure

• Wash and dry hands thoroughly.
• Sterilize intervening "outside" surfaces, such as valves, with 70% alcohol and flame.
• Do not uncap sample tube until immediately before sampling.
• Do not touch inside of tube or cap to any surface.
• If possible, allow cooled, aerated wort to flow directly into tube without using utensils; fill tube about halfway.
• Do not allow sample to overflow onto outside of tube; recap loosely.
• Allow to grow in a warm area (~86°F/30°C) for 3 days.

Notes:

• This test determines whether or not brewing organisms are present without allowing for their identification. Clear, bubble-free wort after 3 days of incubation at room temperature indicates the sample contains no viable brewing organisms. Cloudiness, bubbles or gas escaping when lid is loosened indicates that live organisms are present, which means either your sanitizing regime is inadequate, or sterile materials and surfaces are being exposed and contaminated.

Protocol: Pouring Plates

Equipment

• 5% disinfecting solution
• sterile petri dishes
• sterile prepared media
• permanent marker

Procedure

• Prepare media according to instructions.
• Wash and dry hands thoroughly.
• Clean work area with a 5% disinfecting solution such as Lysol or bleach.
• Lay out, label and date underside of plates and turn each back over, covers up.
• Allow media to cool until it is comfortably-warm to the touch.
• Swirl media occasionally to keep any insoluble components of media suspended.
• Pour 15ml (7ml for small plates) directly into each plate (about 5mm thick), lifting covers just enough to allow for pouring.
• Do not disturb plates until media has solidified.
• Invert and store plates refrigerated in a clean plastic bag.

Notes:

• Pouring media while it is too hot will result in excess condensate inside the plate, which increases the chances of contamination.
• You can check plates for contamination by examining them after 48 hours of incubation in a warm area (~86°F/30°C).
• Wort agar (use in place of commercial nutrient agar) = 30ml house wort, 70ml water, 2g agar; sterilize for 15 minutes at 15psi.

Protocol: Plating Samples Directly

Plated media are generally used to test the purity of liquid samples such as wort, yeast slurry and bottled beer. It is important that the plates do not become contaminated with anything other than the sample you want tested, so store your plates refrigerated and upside-down in a clean plastic box until you are ready to use them.

Equipment

• Starsan or a 5% Lysol or bleach solution
• media plates
• sterile pipettes
• plastic box

Procedure

• Wash and dry hands thoroughly.
• Choose an area that is enclosed or away from drafts and messy or dusty operations.
• Wipe down countertop or table with Starsan or a 5% Lysol or bleach solution.
• Inspect each new plate for contamination; label underside with date, sample name and sample site.
• Liquid sample: place sample on media using a sterile pipette, gently spread over entire surface without gouging media and immediately replace plate lid.
• Filtered sample, see Protocol: Filtering Samples for Plating.
• Swab sample: streak swab tip over entire media, twisting swab to expose all of tip to media surface and immediately replace plate lid.
• Wipe down a plastic box with Starsan or a 5% Lysol or bleach solution.
• Place plates in box upside-down; loosely fit box lid.
• Allow to grow in a warm area (~86°F/30°C) for 3 days. Do not wrap plates or seal plates in any way.

Protocol: Filtering Samples for Plating

If the entire sample is too large to be plated directly (>1.0ml), sample filtration is necessary. The purpose of filtration is to trap any organisms present onto a membrane filter by drawing the sample through it. The filter is then laid directly onto plated media.

Equipment

• Starsan or a 5% Lysol or bleach solution
• 70% alcohol
• gas burner
• vacuum pump
• sterile filtration apparatus
• sterile 0.45 filters
• tweezers
• media plates
• plastic box

Procedure

• Choose an area that is enclosed or away from drafts and messy or dusty operations.
• Wipe down countertop or table with Starsan or a 5% Lysol or bleach solution.
• Wash and dry hands thoroughly.
• Inspect each new plate for contamination; label underside with date, sample name and sample site.
• Dip tweezers in alcohol and light with flame. Allow flame to extinguish on its own.
• Use tweezers to place sterile filter into filtration apparatus; reassemble apparatus.
• Flame mouth of sample vessel and add 100ml to the top receptacle.
• Cap top receptacle and attach vacuum to bottom receptacle. Draw entire sample through filter.
• Sterilize tweezers as above; remove top receptacle and retrieve filter¹.
• Place filter face-up on agar, making sure membrane's entire underside is in contact with media.
• To filter another sample, liberally spray alcohol into apparatus and draw through with vacuum; load with sterile filter.
• Wipe down a plastic box with Starsan or a 5% Lysol or bleach solution.
• Place plates in box upside-down; loosely fit box lid and allow to grow in a warm area (~86°F/30°C) for 3 - 5 days.
• Do not wrap plates or seal plates in any way.

Notes:

• ¹If another sample is to be filtered, assemble apparatus and liberally spray down interior of top receptacle with alcohol; empty using vacuum suction. Sterilize tweezers, put fresh filter in place and repeat filtration procedure.

Protocol: Streak-Plating for Single Colonies / Strain Purification

Equipment

• sterile water (contact-lens saline works well)
• wort agar¹ plate
• yeast culture

Procedure: Streak-Plating for Single Colonies

• Put three drops of sterile water next to each other on the periphery of a wort agar plate.
• Get a film of sample across an inoculating loop (for yeast slurry) or barely touch loop to a colony (for solid culture).
• Dip loop into first water drop and 'wash around until material from loop is suspended in water drop.
• Flame and cool inoculating loop.
• Get film from first drop across loop and transfer to second water drop, washing loop as before.
• Flame and cool inoculating loop.
• Get film from second drop across loop and transfer to third water drop, washing loop as before.
• Flame and cool inoculating loop.
• Get film from third drop across loop and streak across surface of plate, avoiding the three drops.

Procedure: Strain Purification

• Select three or four average-sized colonies which are representative of most of the rest of the colonies.
• Transfer together to wort agar plate and incubate for 2 days in a warm area (~86°F/30°C).
• Wrap plate with parafilm and store upside-down in refrigerator; shelf life is 6 months.

Notes:

• ¹Wort agar = 30ml house wort, 70ml water, 2g agar; sterilize for 15 minutes at 15psi.

Protocol: Viability Testing

Equipment

• microscope with 40x objective
• methylene blue, 0.01% aqueous
• hemacytometer and cover slip, clean and dry

Procedure

• Obtain 2ml of a slightly cloudy suspension of yeast (keep track of your dilutions ¹ if you plan to do a cell count as well).
• Dye sample with 2 drops methylene blue dye.
• Shake sample until color is uniform; allow to sit for 1 minute.
• Place sample in one of the hemacytometer chambers (refer to loading instructions that came with your hemacytometer).
• View under microscope using 40x objective.
• Record numbers of viable (clear) and non-viable (blue) cells² in several same-sized grids and calculate percent viability.

Notes:

• ¹ Confused about making dilutions and calculating dilution factors? Consider a 1:10 dilution. The number on the left side of the colon is the volume (in ml, let's say) you begin with; the number on the right is the volume you end up with. A dense slurry may require a 1:100 dilution, namely, the addition of 1ml yeast to 99ml water,   in order to render it countable. There is another way to get a 1:100 dilution. You may add 1ml yeast to 9ml water, mix, then remove 1ml of that suspension and add it to 9ml of fresh water. In other words, two 1:10 dilutions made in series are equivalent to a single 1:100 dilution. Similarly, three 1:10 dilutions made in series are equivalent to a single 1:1000 dilution. The dilution factor is the product of the numbers on right side of the colon. For example, a 1:10 dilution followed by a 1:4 dilution is equal to a 1:40 dilution and has a dilution factor of 40.

Protocol: Cell Counting

Equipment

• microscope with 40x objective 
• methylene blue, 1% aqueous
• hemacytometer and cover slip, clean and dry  

Staining

• Obtain a few ml of a slightly-cloudy suspension of yeast, keeping track of any dilutions¹ made.
• Dye with methylene blue dye, one drop at a time, until slurry is a cobalt blue.
• Shake sample until color is uniform; allow to sit for 1 minute.
• Place sample in one of the chambers using instructions that came with your hemacytometer.
• View under microscope using 40x objective; blue cells are non-viable, clear are viable.

Counting

• Locate the central square which contains 25 smaller, gridded squares.
• Count the total number of viable yeast in 5 of the squares ² using pattern shown at right.

Calculating Pitch Rate

Notes:

• ¹ Confused about making dilutions and calculating dilution factors? Consider a 1:10 dilution. The number on the left side of the colon is the volume (in ml, let's say) you begin with; the number on the right is the volume you end up with. A dense slurry may require a 1:100 dilution, namely, the addition of 1ml yeast to 99ml water, in order to render it countable. There is another way to get a 1:100 dilution. You may add 1ml yeast to 9ml water, mix, then remove 1ml of that suspension and add it to 9ml of fresh water. In other words, two 1:10 dilutions made in series are equivalent to a single 1:100 dilution. Similarly, three 1:10 dilutions made in series are equivalent to a single 1:1000 dilution. The dilution factor is the product of the numbers on right side of the colon. For example, a 1:10 dilution followed by a 1:4 dilution is equal to a 1:40 dilution and has a dilution factor of 40.
• ² Exclude buds and yeast lying on any two of the four borders of each of the 5 squares.

Protocol: Gram Staining

Equipment

• microscope with 100x objective
• immersion oil
• glass slide
• gram stain kit

Procedure

• Place a drop of water on a clean glass slide.
• Barely touch a flamed and cooled inoculating loop to the colony.
• "Wash" loop in drop of water and evenly smear the suspension around in a pea-sized area; allow to air-dry.
• Hold glass slide about 3 inches above flame and hold for 4 seconds to kill and fix organisms to slide; allow slide to cool.
• Cover smear with crystal violet stain and leave on for 1 minute.
• Rinse and flood slide with iodine mordant; leave on for 3 minutes then drain.
• Slowly drip decolorizing rinse onto smear while holding slide vertically; stop as soon as color ceasing flowing from smear.
• Cover smear with safranin counterstain and leave on for 1 minute.
• Rinse gently with water and allow to air dry.
• Examine under microscope using oil-immersion lens. Gram⁺ is blue or purple; gram^– is red or pink.

Notes:

• The decolorizing rinse is the most critical step. If you continually get gram^– or variable results for known gram⁺ bacteria, you are over-rinsing. Gram⁺ results for gram^– bacteria can result from too dense a smear.

Protocol: Catalase & Oxidase Tests

Equipment

• 3% hydrogen peroxide
• glass slide
• Dryslide oxidase test¹
• absorbent paper

Procedure: Catalase Test

• Drop a little hydrogen peroxide onto the colony in question, or onto bacteria smeared on a clean glass slide.
• A positive result is indicated by the formation of bubbles.

Procedure: Oxidase Test

• Smear a little of the colony in question onto the slide.
• A positive result is indicated by the bacteria turning a very dark purple color within 2 minutes.

Notes:

• ¹ Catalog 3530-75-3 from Difco at (800) 521-0851.

Protocol: Yeast Harvesting & Storage

Equipment

• 70% ethanol or rubbing alcohol
• sanitizer
• storage container(s)
• cold water
• sterile wort

Procedure

• Sterilize valves with 70% alcohol and flame, and sanitize any hoses, utensils and storage containers to be used.
• Catch the middle layer of yeast in a storage container as follows, discarding top and bottom layers:
• CONICAL vessels require draining the bottom trub-filled layer until good color and consistency is

  Seen.

• ROUND vessels require manual harvesting with a paddle (layers mix if allowed to drain through the valve).
• For every gallon of yeast, acidify about 100ml of cold water to ~pH 3 using a food-grade acid.
• For every gallon of yeast, add 1ml¹ or 2ml² sodium chlorite to the acidified water.
• When concoction turns a very pale yellow (at about 15 minutes) add to the yeast slurry, mixing well.
• Allow to sit for a minimum of 30 minutes (longer reaction time ensures effectiveness).
• Use yeast as needed or add an equal volume of sterile, aerated wort and store at 34°F/1°C for up to 2 weeks.
• If stored, test for viability, cell count and contamination before pitching.

Notes:

• It is best to harvest yeast immediately after final gravity has been reached, before diacetyl rest. Even if it is kept very cold yeast should not be stored in the cone or under beer. Finished beer is a nutrient-poor waste product to yeast, and forcing it to reside there will cause the viability to drop precipitously (over 50% within 48 hours), no matter how cold it is kept!
• ¹ DioxyChlor™ from BIRKO at (800) 525-0476
• ² Starzene™ from Five-Star at (800) 782-7019

Table of Brewing Bacteria

Quick Reference

Yeast population increases 4 to 5-fold over entire period in fermentor.

• Repitch 20-25% of yeast mass cropped from last similar-sized batch if viability is >95%.
• Size of starter culture needed for a given brewing capacity:
    ___   bbl capacity   /   10   =   ___   bbl starter culture

sterilizing time vs. temperature

• keep system (not the liquid running through it!) at:
• 160°F (71°C) for 45 minutes
• 170°F (77°C) for 30 minutes
• 180°F (82°C) for 25 minutes

volume conversions

• l = liter
• hl = hectoliter
• bbl = beer barrel
• 3.8 l / gal
• 100 l / hl
• 118 l / bbl
• 31 gal / bbl
• 1.18 hl / bbl
• 0.85 bbl / hl

  °C = 5/9 (°F - 32)

• °F = (9/5 °C) + 32

  ° Plato = % sugar = ¼ (last two digits of specific gravity reading)

• 5° Plato = 5% sugar = 1.020 sg

  OE = original extract in ° Plato

• AE = apparent extract in ° Plato
• RE = real extract, ° Plato
• % alcohol by weight = 0.42 (OE - AE)
• % alcohol by weight = 0.52 (OE - RE)
• % alcohol by weight = 2.22 (RE - AE)
• % alcohol by volume = (% alcohol by weight   X  specific gravity of beer) / 0.791

Glossary

• aerobe: an organism which needs oxygen to survive (obligate aerobes will die without air)
• anaerobe: an organism which does not need air to survive
• aseptic: against or without infection
• contaminant: a relative term referring to any organism which lends undesirable characteristics
• hemacytometer: a glass microscope slide etched with grids of precise dimensions used for counting blood cells
• inoculate: to introduce an organism to a sterile environment, intentionally or not
• morphology: shape or appearance
• petite mutant: a yeast with deficient respiratory abilities, resulting in slow growth and small (petite) colony size
• strains: subspecies or "types" of Saccharomyces cerevisae; most brewing yeasts are strains of these two species
• viability: percent of live cells in a population; not to be confused with vitality, which refers to metabolic activity or vigor of live cells
• wild yeast: undesirable, undomesticated, or non-brewing yeast

© 2001 BSI This handbook is the intellectual property of The Brewing-Science Institute.