QUALITY CONTROL OF LEGUME INOCULANTS WORKSHOP - PROCEDURES SECTION potx

Version date 1/2/2007
Project Number 013/06VIE

Replacing Fertiliser N with Rhizobial
Inoculants for Legumes in Vietnam for
Greater Farm Profitability and Environmental
Benefits



QUALITY CONTROL
OF LEGUME
INOCULANTS
WORKSHOP


PROCEDURES
SECTION



Compiled by:
E. Hartley
G. Gemell
J. Hartley
Australian Legume
Inoculants Research Unit
An NCSI QMS ISO9001:2000
Certified Laboratory 9187-21

1

CONTENTS PAGE

Preparation of sterile Yeast Mannitol Broth (YMB) 2
Autoclave times for liquids 3
Water loss through autoclaving 3
Growth assemblies for test host legumes
Preparing Jensen’s nitrate-free nutrient solution (modified) 5
Plant tubes 6
Gemell Roll Tubes 7
Sub-culturing rhizobial cultures onto agar slopes in McCartney bottles 9
Sub-culturing rhizobial cultures onto agar in Petri plates 10
Gram stain of rhizobial culture 11
Serology (serological agglutination test) 13
Preparing and surface-sterilising seeds with sodium hypochlorite 15
Inoculating YM broths with rhizobia 16
Preparing Yeast Mannitol Agar (YMA) and Congo Red Yeast Mannitol Agar
(CRYMA) 17
Plating out YM broth cultures 19
Measuring the moisture content of rhizobial inoculants 21
Counting rhizobia within an inoculant by Direct Plate count 24
Estimating rhizobial numbers using plant infection and Most Probable
Number (MPN) 26
Isolating rhizobia from nodules 30


2
Ingredients set out before weighing
into a flask containin
g

distilled wate
r

Weighing ingredients
Preparation of sterile Yeast Mannitol
Broth (YMB)

Requirements Reagents for 500mL
• Balance, accurate to 0.01g
• Glass flask or beaker to hold 500mL
• Disposable weigh cups
• Spatula
• 250mL conical flasks with cotton wool
bungs or equivalent
• Autoclavable paper bags
• Autoclave indicator tape
• 0.5 g of yeast extract powder
• 0.1 g of MgSO
4
.7H
2
O
• 0.25 g of K
2
HPO
4

• 5.0 g of Mannitol
• 0.05 g of NaCl
• 500mL of distilled water

Method

1. Pour 500mL of distilled water into a clean glass
flask or beaker
2. Using the balance, weigh out each of the reagents
into separate disposable weigh cups
3. Add the reagents one at a time to the distilled water
and mix thoroughly
4. Carefully pour approximately 50mL of broth into
the 250mL conical flasks. Plug with cotton wool
bungs
5. Place small paper bag over cotton wool bung and
secure with autoclave tape
6. Place autoclave indicator tape on paper bag.
7. Autoclave at 121°C for 20 minutes
8. Wearing protective clothing and heat resistant
gloves, remove flasks from the autoclave
9. Allow to cool before use

3
Sterilization of Single Volumes
0
1000
2000
3000
4000
5000
15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40
Minutes at 121 degrees C

Single volume (ml)
from: Methods in Microbiology 3A p 289. Eds. Norris and Ribbons
Autoclave times for liquids

The graph below shows the autoclave time required when
sterilising single volumes of liquids.



Water loss through autoclaving

It is important to consider the way that diluents are
prepared in terms of accuracy of volumes.
There are differences in accuracy between:
• dispensing water in tubes or bottles before sterilisation
by autoclave
• dispensing sterile water into pre-sterilised tubes or
bottles
If water is dispensed into bottles before autoclaving the
amount of water loss through autoclaving should be taken
into account so that the final volume is correct.
Dispensing sterile water into pre-sterilised bottles has
been found to be more accurate than pre dispensed water
into bottles before autoclaving.
Below is an example of the variation in water loss that
occurs as result of autoclaving. 18.5mL water was

4
measured gravimetrically into 15 McCartney bottles
loosely capped, then sterilised at 121°C for 20 minutes.

The range in water loss was between 2.43 and 7.78%.

Sample
No.
Weight of empty
McCartney's +
lids before
autoclaving (g)
Weight of McCartney's
+ lids with18.5mL
aliquot (g) after
autoclaving
Actual amount of
diluent in
McCartney (mL)
% water
loss
1 55.71 73.76 18.05 2.43
2 53.84 71.62 17.78 3.89
3 51.76 69.53 17.77 3.95
4 52.04 69.74 17.70 4.32
5 54.45 72.12 17.67 4.49
6 53.53 71.18 17.65 4.59
7 52.08 69.69 17.61 4.81
8 53.91 71.50 17.59 4.92
9 51.75 69.31 17.56 5.08
10 54.84 72.37 17.53 5.24
11 55.21 72.73 17.52 5.30
12 52.01 69.48 17.47 5.57
13 52.98 70.36 17.38 6.05

14 56.14 73.42 17.28 6.59
15 54.39 71.45 17.06 7.78



5
Growth assemblies for test host legumes

Preparing Jensen’s nitrate-free nutrient solution
(modified)

A diverse range of legumes of different shapes and sizes
need to be grown in the laboratory.
When growing legumes under controlled environmental
conditions such as a glasshouse or growth cabinet the
plant nutrient solution must contain all the necessary
elements for growth. Jensen's medium provides all
nutrients and trace elements (with the exception of N)
necessary for the growth of most legumes. Growing
legume plants using N-free nutrients will encourage
nodule formation and N
2
fixation.

Jensen’s nitrate-free nutrient (modified) solution

Requirements Reagents
• Electronic balance (accurate to 0.01
g)
• Disposable weigh cups

• Spatula
• Autoclavable glass flask
• Test tubes
• Bromothymol blue pH indicator
• Glass pipette and dispenser
• Heat resistant gloves
• Glass stirring rod

• CaHPO
4
(calcium hydrogen orthophosphate)
• K
2
HPO
4
(di-potassium hydrogen
orthophosphate)
• MgSO
4
.7H
2
O (magnesium sulphate)
• NaCl (sodium chloride)
• FeCl
3
(ferric chloride)
• 1M NaOH (sodium hydroxide)
• Deionised water
• Stock A-Z solution (Gibson) of minor elements
made from: g L

-1

• H
3
BO
3
(boric acid) 2.86
• MnSO
4
.4H
2
O (magnesium sulphate) 2.03
• ZnSO
4
(zinc sulphate) 0.22
• CuSO
4
.5H
2
O (cupric sulphate) 0.08
• Na
2
MoO
4
2H
2
O (sodium molybdate) 0.14
• Bromothymol blue pH indicator solution

6
Notes:

Method

1. Using an electronic balance weigh out:
• 1.0g of CaHPO
4

• 0.2g of K
2
HPO
4

• 0.2g of MgSO
4
.7H
2
O
• 0.2g of NaCl
• 0.1g of FeCl
3


2. Using a measuring cylinder, measure out 1L deionised
water into heat resistant glass flask (or stainless steel
bucket if preparing larger volumes)
3. Add all chemicals to water one at a time in the above
order and mix thoroughly after each addition.

4. Add 1mL of A-Z stock solution
5. Cover flask with aluminium foil and autoclave at 121

°C for 20 minutes (or for the recommended length of
time if a greater volume is being prepared).
6. Wearing protective clothing and heat resistant gloves
remove medium from autoclave.
7. Adjust the pH. to 6.8-7.0.

Plant tubes

This is an enclosed aseptic growth assembly.

Requirements

• Electronic balance
• Weigh cups
• Spatula
• Test tubes and foam or non absorbent cotton wool
bungs (either 18 mm or 25 mm diameter depending on
legume being grown)
• Glass stirring rod
• Autoclave indicator tape
• Autoclavable glass flask / beaker or container
• Agar dispenser
• Wire basket to hold tubes
• Agar
• Jensen's nitrate-free plant nutrient solution

7

Notes:

Method

1. Weigh out 12 g of agar
2. Add the agar to 1L of Jensen's nitrate-free plant
nutrient solution in a container, and stir thoroughly.
3. Place solution into autoclave for the required amount
of time to melt the agar.
4. Carefully remove the molten agar solution from the
autoclave
5. Test and adjust the pH of the medium to pH 6.8-7.0.
6. Using a pre-calibrated dispenser, dispense the required
volume of agar medium into each test tube.
7. Place sponge or cotton wool bung into each tube, and
attach autoclave tape to one of the tube tops or the side
of the container.
8. Place tubes in the autoclave at 121°C for 20 minutes.
9. Wearing protective clothing and heat resistant gloves
remove tubes from autoclave.
10. Before agar has cooled, tip the basket of tubes at an
angle (e.g. lean against a wall) so that the agar within
the tubes sets on a slope.
11. Store in a cool place until required.
NB: When preparing plant tubes to be used for nitrate
controls , follow the above procedure and add 1.0 g KNO
3

L
-1

when testing soybeans, cowpeas, mung beans and 0.5 g
KNO
3
L
-1
for clovers, medics and lucerne.

Gemell roll tubes

This is a modified hydroponic system for growing larger
seeded legumes in an open aseptic growth assembly.

Requirements

• 2L plastic autoclavable jar with screw cap lid
• 18 mm x 15 mm heat resistant test tubes
• Non-bleached absorbent paper towelling
• Measuring jug
• Germinated seedlings
• Forceps
• Absolute alcohol
• Gloves

8
• Jensen’s nitrate-free plant nutrient solution

Method

1. Place the test tube at one end of the paper towel.

2. Wrap the paper towel around the tube leaving about 10
mm overlap at each end of the tube.

3. Tuck the over-lapping paper into the open end of the
test tube, leaving the overlap at the closed end.






4. Place wrapped tube, closed end upwards into the
autoclavable jar.
5. Pour 750-1000mL of plant nutrient solution into the
jar, wetting the towel wrap.
6. Screw lid on loosely, lid and autoclave at 121°C for 60
minutes.
7. Using heat resistant gloves remove assemblies from
autoclave and allow to cool.
8. Wearing gloves, and using pre-sterilised forceps, sow
2 germinating seedlings of the test host plant between
the enclosed end of the tube and the overlapping paper
towel.
9. Replace lids
10. Place assemblies in a controlled environment room at
an appropriate temperature and remove the lids after 2
to 3 days.

9
Sub-culturing rhizobial cultures onto
agar slopes in McCartney bottles

Requirements
• Authenticated source of the rhizobial strain
• Freshly prepared agar slopes in McCartney bottles
• Self adhesive labels
• Marker pen
• Spirit burner
• Inoculation loop
Method

The following steps should be carried out in a Laminar
flow, or a Class II safety cabinet.

1. Carefully write out 2 labels for each of the rhizobial
strains being sub-cultured. Labels should have strain
name, and date of sub-culture clearly written.
2. Place labels on the lids and side of the McCartney
slopes, and place next to original source of rhizobia to
be sub-cultured.
3. Sterilise the inoculation loop by holding it in a burner
flame, starting at the top of the wire and slowly

moving down to the loop until the loop glows red.
4. Aseptically unscrew the McCartney slope lid
containing the original culture (re-check to make sure
that the labels are correct), pass mouth of bottle
through the flame. Cool the loop in the agar before
touching the rhizobial culture.
5. Take a loop-full of culture, and quickly pass mouth of
bottle through the flame before replacing the lid
6. Ensuring that the inoculation loop does not come into
contact with any surface, remove the lid from the fresh
McCartney slope, pass mouth of bottle through the
flame, and place the inoculation loop with the rhizobia
onto the surface of the agar slope. Spread the loop-full
of rhizobia across the agar surface taking care not to
touch the sides of the McCartney bottle.
7. Remove the inoculation loop, and pass the mouth of
the bottle through the flame before replacing the lid.
8. Re-sterilise the inoculation loop.
9. Place McCartney slopes upright in a container, and
incubate at 26°C for the required length of time.

Ensure the loop is flamed full
length
Subculture onto YMA slope

10
Notes:
The desired results of each of
these procedures, is that less of
the initial suspension is

distributed over each consecutive
streak, resulting in isolated single
colonies in the last 2 quadrants.

Sub-culturing rhizobial cultures onto
agar in Petri plates

Requirements

• Rhizobial culture to be examined
• Freshly prepared YMA and CRYMA plates
• Marker pen
• Spirit burner
• Inoculation loop

Method

The following steps should be carried out in a Laminar
flow or Class II safety cabinet.

1. Label and date the underside of the Petri plates with
the rhizobial strain name, and date of sub-culture.
2. Keep the Petri plates inverted.
3. Sterilise the inoculation loop by holding it in a burner
flame, starting at the top of the wire and slowly
moving down the loop, until the loop glows red. Cool
the loop by placing it on the surface of the agar.
4. Take a loopful of the rhizobia suspension to be
cultured, lift the agar plate with the other hand (leaving
the lid face up on the work surface of the cabinet);

streak the loopful of culture across the surface of the
agar backwards and forwards several times, at the top
of the plate.
5. Turn the loop over, and using the other side continue
to spread the rhizobia suspension over the second
quadrant of the plate, at right angles to the initial
streak. Place the agar plate back into the lid. Re-
sterilise the inoculation loop.
6. Pick up agar plate and touch the loop into the agar at
the edge of the plate to cool it. Beginning at the edge
of the second quadrant of the streak, make 4 single
streaks into the 3rd quadrant of the plate.
7. Turn the loop, turn the plate, and using the clean side
of the loop, barely crossing the edges of the previous 4
streaks, do a final zig-zag streak in the 4th quadrant of
the plate and replace agar plate into the lid.
8. Re-sterilise loop.
9. Keep agar plates inverted and incubate at 26°C for the
required length of time.
Streak of rhizobia on CRYMA

11
Notes:

Gram stain of rhizobial culture

Requirements

• Crystal violet solution
• Iodine concentrated solution

• Dilute iodine (iodinated alcohol) solution
• Safranin solution
• Cleaned glass microscope slide
• Glass marker (wax pencil)
• Sterile water
• Spirit burner
• Access to wash water at a sink
• Slide rack
• Paper towel
• Microscope with oil immersion lens (100x)
• Immersion oil
• Timer or watch

Method

1. Clean a glass microscope slide by soaking in ethanol
and wiping dry.
2. Label the slide using the wax pen. Draw a small circle
for each smear of cells.
3. Add a small droplet of sterile water to the clean slide
within the circle.
4. Using a burner, sterilise an inoculation loop.
5. Collect a small loop of culture from the sample and
evenly mix it in the small drop of sterile water on the
slide.
6. Allow the smear to air-dry then heat fix by quickly
passing the slide through a burner flame a few times.

To be carried out on a slide rack over a sink with access
to cold water

7. Cover the smear with a drop of crystal violet and leave
for 1 minute.
8. Wash gently with cold water for 5 seconds.
9. Cover the smear with one drop of concentrated iodine
solution and leave for 1 minute.
10. Pour off concentrated iodine and cover smear with
iodinated alcohol and leave for 5 minutes.
11. Wash gently with water for 5 seconds
.
Add Safranin to counterstain cells

12
Notes:

12. Counterstain the smear with one drop of Safranin and
leave for 3 minutes.
13. Wash smear gently with water, allow to air-dry.

Microscopic examination of Gram stain

1. Add a drop of emersion oil onto stained smear, and
examine under a microscope at a magnification of
x1000 using a 100x oil emersion lens
2. If slide shows any Gram +ve cells or Gram -ve cells
which are not bacilli, the culture is not a rhizobia
3. If slide shows Gram -ve bacilli alike in size and shape,
the culture fits the criteria for rhizobia

N.B. Time of application of each stain should be accurate,
as leaving any stain on for either too long or too short a

time may affect the results.


Serology (serological agglutination test)

Requirements

• Rack containing metal capped test tube
• Pipette calibrated to dispense aliquots equal to 16
drops
• Sterile plugged pipette tips
• Vortex mixer
• 3 serology tubes and rack
• Water bath at 100°C or similar apparatus
• Water proof marker pen
• Spirit burner
• Inoculation loop
• Sterile Pasteur pipette
• 0.85% NaCl
• 5mL glass pipette
• Pure culture of strain to be tested
• Frozen antisera of strain to be tested at appropriate
titre (1:20 dilution). Do not use concentrated antisera.

Method

To be carried out in a lamina flow safety cabinet if
possible.
Step 1
1. Using a waterproof marker pen, label a metal–capped

test tube with the batch number and strain being tested
Examine the Gram stained slide

13
2. Using a glass pipette and dispenser, dispense 3.5mL of
0.85% NaCl (saline) into the tube
3. Pre-sterilise the inoculation loop, by placing in a
flame, and allow to cool before taking a loopful of
pure culture and adding it to the tubes containing NaCl
4. Mix solution using the Vortex mixer and place tubes in
the water bath set at 100°C and boil for 30 minutes to
inactivate flagella `H' antigens.
Step 2
1. Remove antiserum (1:20 dilution) of the nominated
strain from the freezer. Allow it to thaw before use.
2. Using heat resistant gloves, remove test tube from
water bath. Allow to cool in a rack.
3. Using water proof marker pen, label 3 serology tubes
with strain names and the third tube labelled # as the
control
4. Dispense 16 drops of the boiled suspension into the
three serology tubes.
5. Dispense 2 drops of a 1:20 dilution of antiserum of the
nominated strain in the first 2 serology tubes
6. Dispense 2 drops of 0.85% saline into the 3rd serology
tube labelled # (control).
7. Leave serology tubes at room temperature (warm)
overnight
8. After 12h, check suspension for cell agglutination.

14
Notes:

Preparing and surface-sterilising seeds
with sodium hypochlorite

Requirements

• Seeds - to be surface sterilised
• Pyrex conical flask and rubber bung
• Stopwatch or timer
• Protective clothing
• Gloves
• “Miltons” tablets as a source of sodium hypochlorite
• Absolute alcohol
• Sterile water
• Sink with access to running water

Using sodium hypochlorite

1. Place sufficient amount of seeds in a clean conical
flask stoppered with a rubber bung.
2. Add absolute alcohol to cover the seeds and wash by
gentle agitation for a few seconds then immediately
pour off the excess into the sink.
3. Pour sufficient bleach (NaOCl) into flask to cover
seeds and rotate the flask to wet all the internal area of
the flask including the bung.
4. Set stopwatch and leave seeds soaking for 4 minutes.
5. Pour off the bleach carefully into sink with running

water, keep the seeds trapped inside the flask.
6. Using cold sterilised water, cover the seeds and wash
carefully.
7. Tip the water away using the bung to stop the seeds
from being washed out of the flask. Repeat the rinse
procedure a total of 9 times.
8. After the final 9
th
rinse, leave seeds soaking in sterile
water until the seeds fully imbibe except for soybeans
which are drained immediately
Pour off excess li
q
uids between ste
p
s

15
Notes:

Inoculating YM broths with rhizobia

The following steps should be carried out in a Laminar
flow, or a Class II safety cabinet.

Requirements


• Pure culture of rhizobia
• Sterile YM broth

• Marker pen
• Inoculating loop
• Spirit Burner


1. Carefully label side of flask and the paper cap. Labels
should have strain name, and date of inoculation and
operators name clearly written.
2. Sterilise the inoculation loop by holding it in a burner
flame, starting at the top of the wire and slowly
moving down to the loop until the loop glows red.
3. Aseptically unscrew the McCartney slope lid
containing the original culture, pass mouth of bottle
through the flame. Cool the loop in the agar before
touching the rhizobia culture.
4. Take a loop-full of culture, and quickly pass mouth of
bottle through the flame before replacing the lid.
5. Ensuring that the inoculation loop does not come into
contact with any surface, remove the bag and cotton
wool bung from the YM broth, pass mouth of flask
through the flame, and place the inoculation loop with
the rhizobia into the broth and wash the cells off the
loop.
6. Flame the top of the flask before replacing the cotton
wool bung. (Care must be taken not to wet the bung)
7. Sterilise the inoculation loop.
8. Gently mix the broth, and place in an incubator shaker
at 26°C -28°C until broth becomes milky in colour. (3-
7 days depending upon the rhizobia being cultured)

Ensure the loop is flamed full
length

16
Preparing Yeast Mannitol Agar (YMA)
and Congo Red Yeast Mannitol Agar
(CRYMA)

For 1 litre of agar

Requirements Reagents
• Electronic balance accurate to 0.01g
• Heat resistant 2L glass flask
• Disposable weigh cups
• Spatula
• 1L measuring cylinder
• Sterile Petri plates
• Clean work station
• Heat resistant gloves
• Aluminium foil
• Autoclave indicator tape
• Sterile glass pipette and pipette dispenser
•
• yeast extract powder
• MgSO
4
.7H
2

O (magnesium sulphate)
• K
2
HPO
4
(dipotassium hydrogen
orthophosphate)
• mannitol
• agar
• NaCl (sodium chloride)
• congo red indicator*
(1:400 solution (1.0 g of congo red
indicator to 400mL d H
2
O)
• 70% alcohol
• 1 litre distilled water
• NaOH (sodium hydroxide)
• pH indicator




Method

1. On the balance weigh out each ingredient into separate
weigh cups using the spatula:
• 1.0 g of yeast extract
• 0.2 g of MgSO
4

.7H
2
O
• 0.5 g of K
2
HPO
4

• 0.1 g of NaCl
• 10.0 g of mannitol
• 15.0 g of agar

If preparing YMA with added dye prepare
• 10.0mL of congo red*
2. Measure out 1L of distilled water
3. Empty the 1L of distilled water into a 2L heat resistant
glass flask.
4. Add each of the above chemicals one at a time,
mixing the solution thoroughly after each chemical
has been added.
Weighing yeast powder
Ingredients set out before weighing into
a flask containin
g
distilled wate
r

Pour 10mL congo red into a measuring
c
y

linder read
y
for use

17
Notes:

*Only add congo red if preparing Congo Red YMA i.e.
CRYMA; do not add if preparing YMA.
5. Seal flask with aluminium foil and attach autoclave
heat indicator tape on top of the foil.
6. Sterilise agar in autoclave at 121°

C for 20 minutes (or
for the required length of time if a greater volume is
being prepared).
7. Wearing protective clothing and heat resistant gloves,
remove the molten agar medium from autoclave when
sterilising cycle has completed.
8. If necessary, adjust the pH to 6.8 - 7.0 by adding 1M.
NaOH (approx. 0.1mL of NaOH per 1L of medium).
9. Allow the molten agar to cool in a separate area to
approximately 50 °C, ready for pouring into Petri
plates.

Pouring plates

The following procedures should be carried out in either a
Laminar Flow or if available, a Class II Safety cabinet.
The cabinet should be pre-sterilised by UV light 5-10

minutes before use with the fan-forced clean air flowing.
If these are not available, pre-sterilise a work station with
70% alcohol before use.

1. Remove sufficient Petri plates from sterile wrapping
and place in stacks of 5 or 10.
2. When agar has cooled to approx. 50°C, dispense about
25mL of agar into each 90 mm plastic pre-sterilised
Petri plate.
3. Allow agar to set, and then invert plates and store in a
cool place until required.







Try to have plates ready 24
hours before use

18
Plating out YM broth cultures

Requirements

• A rack with sufficient number of sterile Eppendorf
tubes for a complete serial dilution to the level
required.
• Micro Pipette, range 50-250µL

• Pre-sterilised plugged micro pipette tips (250µL)
• Vortex mixer
• Spirit burner
• Glass beaker
• Glass spreader
• Disposal container
• Marker pen
• Aseptically prepared Petri plates containing YMA
and CRYMA, pre-poured at least 24 hours before
use. (need 2 plates for each dilution level to be
counted)
• 100mL of absolute alcohol

Method

1. Pipette 0.1mL (100µL) aliquot from the broth culture
and aseptically transfer it to the first Eppendorf tube
of the dilution series containing 0.9mL of water
diluent. Close Eppendorf lid.
2. Mix contents well by placing the Eppendorf tube on a
vortex mixer for 2-3 seconds. Discard pipette tip.
3. Repeat the above procedure, taking a 0.1mL aliquot
from the first Eppendorf tube and adding it to the next
tube in the dilution series, then repeat again until the
desired dilution levels have been reached, e.g. 10
-6
and
10
-7
for a broth culture that is approximately 10

9
cells
per mL.
4. Take a 0.1mL aliquot from the appropriate dilution
levels to be counted and dispense it onto the agar
surface of the paired plates labelled for that dilution.
5. Sterilise the glass spreader by dipping into the beaker
containing absolute alcohol, and passing it through a
burner flame.
6. Allow a few seconds for the alcohol to burn off and
the spreader to cool, then spread the 0.1mL aliquot
sample evenly across the surface of the agar ensuring

19
that neither the spreader nor sample touches the sides
of the Petri plate.
7. Place inverted Petri plates into an incubator set at 26

°
C for the required time. This depends upon the type
of rhizobia being counted and is 3-5 days for the “fast-
growing” strains, and 5-10 days for “slow-growing”
rhizobia.
Using the flamed glass spreader evenly
spread the 0.1mL aliquot over the agar
surface.


20
Perform QC Tests on Commercial

Inoculants

Measuring the moisture content of
rhizobial inoculants

One of the principal factors affecting survival of rhizobia
in a legume inoculant is the moisture potential of the
inoculant. Basically, the inoculant requires an amount of
moisture to compensate for losses during the expected
shelf life. High moisture content within a carrier has little
effect on the survival of rhizobia, but can encourage the
growth of other micro-organisms which may inhibit the
growth and survival of the rhizobia. On the other hand, if
the moisture content of the inoculant is too low, the
rhizobia become stressed resulting in death.
Therefore, checking the moisture potential of inoculants is
one of the key tests in quality evaluation.

Requirements

• Electronic balance
• Glass Petri plates
• Marking pen
• Peel-off sticky labels
• Calculator or data spreadsheet
• Hot air oven pre-calibrated to 80°C
• Heat resistant gloves

Method

1. Place a peel-off sticky label to the lid of a glass Petri
plate.
2. Weigh glass Petri plate and label. Record the weight
on the label.
3. Place approximately 10 g of inoculant sample into the
glass Petri plate.
4. Re-weigh Petri plate and sample. Record this weight
on the label beneath the previous entry.
5. Place Petri plate in a hot air oven set at 80°C. Leave
for 48 h.
6. Using heat resistant gloves remove Petri plate from hot
air oven and allow it to cool. When cool, weigh the
Petri plate plus dry inoculant. Record weight onto
label beneath other two entries.
Weighing oven-dried peat

21
Moisture Potential Curve of Carrier "a"
0
1
2
3
4
5
6
7
8
20 40 60 80 100 120 140 160 180 200 220 240 260
% Mo istur e
-Log Pa

Moisture Potential Curve of Carrier "b"
2.5
3
3.5
4
4.5
5
5.5
6
6.5
7
30 40 50 60 70 80 90 100 110 120 130 140
% Mois tur e
- Log Pa
Calculation of results

(i) Calculate the % moisture content of the peat
sample (wet weight basis):

Remove the peel-off labels and calculate the moisture
potential of the sample as shown in the following example.

Weight of empty Petri plate (a) = 49.6 g
Weight of Petri plate + wet inoculant (b) = 68.5 g
Weight of Petri plate + dry inoculant (c) = 58.0 g

b-c x 100 = 68.5 - 58.0 x 100
b-a 68.5 - 49.6

= 10.5

x 100
18.9

= 55.5%

The moisture content of the sample is 55%.

(ii)Calculate the moisture potential of the sample:

Calculate the percent moisture of the inoculant on a
dry weight basis. (the ratio of water to peat in the
sample).


b-c x 100 = 68.5 - 58.0 x 100
c-a 58.0 - 49.6

= 10.5
x 100
8.4

= 125.0%


The weight (volume) of water in the example inoculant
represents 125% more than the weight of the peat.
From the predetermined moisture potential curves
prepared specifically for the 2 inoculant carriers,
determine the moisture potential of your sample.
Note how they differ between carriers.

22
NB. All inoculant carriers have a different moisture
potential, therefore before a result can be determined for
that specific carrier, a moisture potential curve must have
previously been prepared.

Moisture curves for two carriers shown on the previous
page illustrate the variability in the amount of moisture
required to achieve similar moisture potentials.


Counting rhizobia within an inoculant
by Direct Plate count

Set up the equipment as for “plate out YM broth
procedures”

1. Surface sterilise the corner of the packet and cut the
corner off with a pair of sterilised scissors.
2. Weigh 10g of inoculant into a labelled sterile plastic
bag.
3. Add by weight or measure an aliquot of 90mL of
sterile water to bag. (This will give an initial dilution
of 1 in 10)
4. Blend peat and water for 30 seconds.
5. Using the 50-250 µL Pipette and a 250µL plugged
pipette tip, take 100 µL( 0.1mL) aliquot from the
initial sample in the bag and place into the first

Eppendorf tube of the dilution series containing 0.9mL
of water diluent. Close Eppendorf lid.
6. Mix contents well by placing the Eppendorf tube on a
vortex mixer for 2-3 seconds. Discard pipette tip.
7. Repeat the above procedure, taking a 0.1mL aliquot
from the first Eppendorf tube and adding to the next
tube in the dilution series, then repeat again until the
desired dilution levels have been reached, (e.g. 10
-6

and 10
-7
for peat inoculants).
Cut across the corner of the surface-
sterilised peat inoculant packet with a
pair of sterile scissors

23
8. Take a 0.1mL aliquot from the appropriate dilution
levels to be counted and dispense it onto the agar
surface of the paired plates labelled for that dilution.

9. Sterilise the glass spreader by dipping into the beaker
containing absolute alcohol, and passing it through a
burner flame.







10. Allow a few seconds for the alcohol to burn off and
the spreader to cool, then spread the 0.1mL aliquot
sample evenly across the surface of the agar ensuring
that neither the spreader nor sample touches the sides
of the Petri plate.
11. Place inverted Petri plates into an incubator at 26
°
C
for the required time. This depends upon the type of
rhizobia being counted and is 3-5 days for the “fast-
growing” strains, and 5-10 days for “slow-growing”
rhizobia.
NB: When counting samples for research purposes, repeat
procedures 1-6 at least 3 times to provide replicate counts
of the test sample. For this workshop you will only be
doing 2 replicate counts.
Using the flamed glass spreader, evenly
spread the 0.1mL aliquot over the agar
Dispensing 0.1mL dilution onto

CRYMA plate

24
Inoculation of plants in test tubes from a
serial dilution using 1mL aliquots.
Estimating rhizobial numbers using
plant infection and Most Probable
Number (MPN)

The number of viable rhizobia in inoculants and rhizobial
products can be estimated by (MPN) plant infection tests.
This method of counting rhizobia can be used in
conjunction with, or instead of, direct plate counts where
there are likely to be high numbers of other micro-
organisms in the sample, making identity and counting of
the rhizobia impossible. The plant infection method of
counting depends on the ability of specific rhizobia to
produce nodules on a chosen species of legume, and
assumes that a single rhizobial cell added to a test-plant is
sufficient to cause nodulation. The estimate obtained from
this test is known as the 'Most Probable Number' (MPN)
of rhizobia in a test sample.

With the method, the number of test-plants with positive
nodulation at specific dilution levels is the basis for
determining rhizobial numbers in the test samples. For
routine laboratory counting, 3 test-plants per dilution level
are used.

This procedure is normally done in conjunction with the

previous procedure “Counting Rhizobia within an
Inoculant by Direct Plate Count”.


1. Using the 250-500 µL pipette and a 500 µL plugged
pipette tip, take 0.1mL aliquot from the initial sample
and place into first Eppendorf tube of the dilution
series containing 0.9mL. Mix well by placing the
Eppendorf tube on a vortex mixer for 5 seconds.
Discard pipette tip.
2. Repeat the above procedure until the desired dilution
levels have been reached. The top dilution levels
should be estimated to give positive nodulation of
seedlings at the initial dilution levels and negative
nodulation of seedlings at the highest dilution level
(e.g. positive nodulation at 10
-7
and 10
-8
levels and
negative nodulation at 10
-9
or 10
-10
for most peat
inoculants).
3. To inoculate the plants at the correct dilution levels to
be counted, e.g. 10
-7
, take 3 x 0.1mL aliquots from the

10
-6
and add to the 3 plant tubes labelled for 10
-7

dilution level. Begin dispensing from the highest
dilution level (i.e.10
-10
), then work backwards to the
lowest dilution level. This enables 1 pipette tip to be
used for the complete dilution series and prevents