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Laboratory manual for Advanced Animal nutrition Research

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School of Animal and Range Sciences
Laboratory Manual for Advanced Animal utrition Research
BY:
Merga Bayssa (Ph.D., Animal nutrition)
Tadesse Bokore (M.Sc., Animal Production)
Feleke Tigabieneh (B.Sc. in Chemistry)
Hawassa University,
Hawassa, Ethiopia
December, 2022


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PREFACE
This laboratory manual represents an attempt to summarize and consolidate a
considerable amount of information relative to laboratory procedures and experimental
techniques that are used commonly in animal nutrition laboratory of the School of
Animal and Range Sciences. It was originally designed to support practical courses of
advanced animal nutrition, Ruminant nutrition and Feeding systems, Monogastric
nutrition, Dairy animal nutrition and Feed stuff evaluation at School of Animal and
Range Sciences College of Agriculture, Hawassa University and attempts have been
made to include basic safety rule and regulation as well as analytical procedures. As
such, the manual served the school for practical courses for more than 8 years as routine
procedures for both under graduate and graduate programmes. Basic inputs have been
included from outlines of the instructors' course notes to improve the contents of the
manual. The late Dr. Aster Abebe deserves special recognition for their immense
contributions on manual to be used as routine laboratory exercise for animal nutrition
courses.
Basically, the text is composed of routine procedures of the main activities in the
laboratory with available resources such as laboratory safety rules and regulations,
Processing and storage of biological samples, proximate analysis, in vitro digestibility
and gas production, spectrophotometric method of phenolics and mineral analysis,
complexometric titration methods, measurement of methane and other gases of rumen
fermentation, In sacco degradability of feed stuffs and conventional in vivo digestibility
and metabolic trials and growth experiments. In addition, measurements of silage quality,
use of markers in nutrition studies, and microbiology of ruminants were included in the
manual. It is not intended to be complete in every detail, and outside reading by students,
scholars and experts will often be necessary for improvement in the quality of the
manual. The authors hope that this manual will serve as a useful reference in the years to
come for those students who select experimental animal nutrition as a career.
Merga Bayssa (Ph.D.)
Tadesse Bokore (M.Sc.)
Feleke Tigabieneh (B.Sc.)


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Table of Contents
Contents page
Preface ----------------------------------------------------------------------------------------- 2
Practical 1: Orientation of standard laboratory safety rules and regulations…….....5
Practical 2: Sample collection and preparation for chemical analysis………...….15
Practical 3: Writing laboratory reports as scientific paper ………………………...18
Practical 4: Determination of moisture content…………………………………….20
Practical 5: Determination of absolute dry matter and ash………………………....22
Practical 6: Determination of crude fat (Ether extract) …………………………….26
Practical 7: Determination of crude fiber content……………………………..…….29
Practical 8: Determination of crude protein………………………………………....31
Practical 9: The detergent system of Fiber analysis………………………………….34
Practical 10: Determination of Urea………………………………………………….38
Practical 11: Determination of Uric Acid…………………………………………….40
Practical 12: Determination of In vitro Digestibility…………………………………44
Practical 13: Determination of In sacco (Nylon bag) degradability………………….49
Practical 14: Determination of In vitro gas production………………………………..51
Practical 15: Conventional digestion/metabolic experiments………………………….57
Practical 16: Determination of Incriminating substance in feeds………………………64

Laboratory manual for Advanced Animal nutrition Research 4
Practical 17: Determination of Major minerals in feeds and biological samples………78
References……………………………………………………………………………....


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Practical 1: Orientation of standard laboratory safety rules and regulations
One of the major concerns of any worker is to be safe while on the job. Both from the
aspect of personal safety and from the aspect of liability for employees, the concern is
justified. Most areas of the laboratory activities involve certain potential hazards and our
analytical laboratory is certainly no exception. The key to job safety in spite of this is the
recognition of the hazards involved in laboratory work, an understanding of what can be
done to reduce the risk of having an accident, and knowing the proper responses to
accidents that may occur.
The following list contains the most common hazards and types of dangerous materials
associated with working in a laboratory which handles chemicals and feed samples, and
includes some suggestions for steps which may be taken to help prevent an accident from
occurring. The list does not include all possible situations which may arise; many types
of hazards may be specific to a particular facility or type of process.
1. Poisons:
Many of the chemicals commonly used in the lab are deadly poisons. Some of these such
as carbon tetrachloride or mercury can be absorbed through the skin and may build up
over a long period of time to dangerous levels. Others such as cyanide may take on a
gaseous form that is extremely dangerous when breathed in. Warning labels on chemicals
used should be read and understood. If the chemical being used is poisonous, special care
should be taken to assure that the material will not be ingested or absorbed through the
skin. All such reagents should be clearly labeled as to its poisonous nature.
2. Explosive Materials:
Almost all labs use acetone, azide compounds, and many other explosive chemicals.
Other chemicals which may not be explosive alone may form explosive compounds with
other non-explosive chemicals. Heat, an electric spark, sudden shock, pressure, or even
contact with air may trigger an explosion from some compounds. Whenever explosive
solvents such as ether or acetone are being used in the lab, open flames must not be used.


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Procedures using these chemicals should be carried out in the fume hood, if possible,
with the fan on. If sample digestions involve the use of perchloric acid, an explosion
proof fume hood rated for perchloric acid must be used. Analytical procedures must be
followed exactly as written using the chemicals specified. Substitutions of chemicals or
alterations in the procedures may cause dangerous reactions to occur.
3. Electrical Shock:
This usually occurs due to improper grounding of instrumentation or improper contact
between the analyst, electricity, and water. Make sure that any instrument is grounded
before use. Avoid using electrical instrumentation near sinks or other sources of water.
Do not operate electrical instruments while standing in water. If an instrument does get
wet do not use it until it has been dried out and has been determined safe for use. Do not
have electrical outlets placed near sinks. All permanent wiring should be installed by a
qualified electrician. Do not overload electrical circuits in the lab. Know the location of
circuit breaker boxes that control circuits in the lab and have the breakers clearly
marked.
4. Mechanical hazards:
High speed centrifuges, sharp equipments, laboratory grinding mills, feed processing
mills, meat mincer(chopper) and vortex mixers(blenders) should not opened while they
are running and should be handled carefully to avoid potential mechanical dangers on the
users.
5. Toxic Fumes:
These are generated as part of many routine procedures. An example of this is the
generation of sulfur trioxide fumes during the analysis for total Kjeldahl nitrogen. This
becomes dangerous when a properly operating fume hood is not used. Observe
precautions printed on all reagent bottles. If the analyst uses a chemical which emits toxic
fumes the work must be done in a fume hood. The fume hood must be inspected at least
once each year to assure an adequate air displacement and to check for leaks in the duct


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work. Spills of such materials, such as mercury, must be cleaned up immediately using
appropriate procedures.
6. Corrosive Materials:
Most labs use concentrated acids and bases for a wide variety of purposes. These not only
are corrosive to laboratory equipment and instrumentation, but can also damage clothing
and cause severe burns. This is especially critical when these materials come in contact
with the eyes. Concentrations of acids and bases should always be specified on the label.
When making up dilutions of acids always add the acid to the water not water on acid
otherwise violent splashing or explosion will occur. Make such solutions cautiously and
slowly, expecting the solution to get very hot. Quantities of these materials of one liter or
more are best stored in unbreakable containers. Put together a kit to handle spills of acids
and bases and keep this in a handy location. Always wear eye protection, apron, and
gloves when handling concentrated acids or bases.
7. Fire:
Fires are usually caused by improper handling of chemicals or from overloaded or
improper electrical conditions. Follow proper storage procedures for all reagents. Dispose
of chemicals in a safe manner. Observe shelf lives of any reagents which are so dated.
Use common sense when using open flames. Know the service capacity of the electrical
circuits in the lab to avoid creating an overloaded condition. Label all circuit breakers
according to major equipment operated on each circuit.
A. General Lab Considerations:
1. Cylinders of compressed gases are extremely dangerous and require special precautions
for moving and storage. If the valve is knocked off accidentally the cylinder may be
propelled with rocket force, damaging almost anything in its path. When moving
cylinders the valve protection cap must be installed and the cylinder should be strapped to
a trussed handcart. For storage and for use, the cylinders should be chained or strapped
securely to prevent them from being knocked over.


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2. Chemicals and reagents should be stored in an adequate storeroom. Heavy items should
be stored as near as possible to the floor. All chemicals should be clearly labeled and
dated. The storage room should be properly ventilated to prevent a possible buildup of
vapors or heat. Care should be taken to assure that incompatible materials are not stored
together.
List of chemicals and their contraindication during storage are given.
3. The lab must have at least one emergency eye wash and shower. These must be
inspected and flushed at least once per month.
4. The location of fire extinguishers, fire alarms, and telephones must be clearly visible.
An emergency telephone number list should be developed and posted near the telephone.
5. A first aid kit must be readily accessible in case of emergency. Proper disposal
procedures should be followed for any outdated or spent reagents.
6. The local fire department may offer information or assistance in disposing of hazardous
chemicals. Broken glass and glass containers should be disposed of in a container
designated for only this type of wastes
7. Response to Emergencies:
In many types of emergencies quick response may mean the difference between having a
close call or having a disaster. One of the best ways to assure a quick response to
emergencies is to make sure that laboratory personnel are adequately trained in the use of
safety equipment and in first aid procedures. Safety equipment training should include
the use of fire extinguishers, emergency shower and eyewash, respirators, and all
other safety equipment which would be appropriate for the particular facility. First aid
training should include basic first aid as well as a course in cardiopulmonary
resuscitation.
In summary, the best way to prevent accidents from happening is to know the procedures
and materials that must be used, to understand the hazards associated with them, and to
use the proper safety precautions and equipment. The best way to prepare laboratory
personnel to react to an emergency situation is by providing the necessary safety
equipment and by providing training in their use.


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B. LABELIG
Adequate labeling of containers of laboratory reagents is essential to providing a safe
working environment in any laboratory. Ethiopian federal government law specifies that
Identity labels, showing contents of containers (including waste receptacles) and
associated hazards” are required. These also state that “employers shall ensure that labels
on incoming containers of hazardous chemicals are not removed or defaced.
It is recommended that labels on containers of chemicals acquired by the laboratory
should include the following information:
A product name, trade name, chemical name or generic name if the product or trade name
is used,
A signal word to draw attention and designate the degree of hazard such as:
1) DAGER shall mean most serious hazard
2) WARIG shall mean a lesser hazard
3) CAUTIO shall mean the least hazard,
c) A statement indicating the level of hazards occur with customary use or handling of
the substance, for example causes burns or vapor hazardous.
d) Date of preparation and / or expiration.
Example: The label for a 1 Normal Sulfuric Acid solution would be as follows:
Sulfuric Acid
H2SO4
1N
Danger – Causes Burns
Prepared: 01/10/2015


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Several labeling tools are available, and each has its place in the laboratory. Most beakers
and flasks will have a hexagonal space of ground glass which can be written on to
identify it. A lead pencil should be used for this type of marking. Grease pencils are
primarily used for temporary labeling. It should be noted that the grease pencil marking
will readily rub off. Commercially available labeling tape is especially useful in many
situations. It may be purchased in several different colors, and may be blank or imprinted
with a form which may be filled out to provide the necessary information. High
temperature markers are available for marking on surfaces that are exposed to extreme
high temperature environments, such as Gooch crucibles. The marks become permanent
after heat is applied. Whatever labeling techniques you use, be consistent, and remember
that the label is intended not only for convenience but also for safety.
Table: Standard and Safe Chemical storages.
These chemicals Should not be stored with:
Acetic acid
Chromic acid, nitric acid, hydroxyl compounds, ethylene
glycol, perchloric acid, peroxides, permanganates
Acetylene Chlorine, bromine, copper, fluorine, silver
Ammonium nitrate Acids, powered metals, flammable liquids, chlorates, nitrites,
sulfur, finely divided organic or combustible materials
Carbon, activated Calcium hypochlorite, all oxidizing agents
Chlorates Ammonium salts, acids, powdered metals, sulfur, finely
divided organic or combustible materials
Chromic acid Acetic acid, naphthalene, camphor, glycerine,
turpentine,alcohol, flammable liquids in general
Chlorine Ammonia, acetylene, butadiene, butane, methane, propane (or
other petroleum gases), hydrogen, sodium carbide, turpentine,
benzene, finely divided metals
Copper Acetylene, hydrogen peroxide
Flammable liquids Ammonium nitrate, chromic acid, hydrogen peroxide, nitric
acid, sodium peroxide, the halogens


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Hydrocarbons Fluorine, chlorine, bromine, chromic acid, sodium peroxide
Hydrofluoric acid,
anhydrous
Ammonia, aqueous or anhydrous
Hydrogen peroxide Copper, chromium, iron, most metals or their salts, alcohols,
acetone, organic materials, aniline, nitromethane, flammable
liquids, combustible materials
Hydrogen sulfide Fuming nitric acid, oxidizing gases
Mercury Acetylene, fulminic acid, ammonia, oxalic acid
Nitric acid,
concentrated
Acetic acid, aniline, chromic acid, hydrocyanic acid, hydrogen
sulfide, flammable liquids, flammable gases
Oxalic acid Silver, mercury
Potassium
permanganate
Glycerin, ethylene glycol, benzaldehyde, sulfuric acid
Silver Acetylene, oxalic acid, tartaric acid, ammonium compounds
Sulfuric acid Potassium chlorate, potassium perchlorate, potassium
permanganate, or similar compounds with light metals
Routine lab safety rules in nutrition laboratory
1. Laboratory eatness: Clean and neat work areas avoid risk of damage to clothing and
books and injury from spilled chemicals. Neatness also reduces fire hazard.
2. Laboratory Conduct: Fooling around in the laboratory can be hazardous.
3. Working with Glassware: Remove frozen glass stoppers with proper equipment.
Broken or chipped glassware should be discarded. Properly support glassware with ring-
stands and clamps when heating and use cork rings with round-bottom flasks.
4. Working with Glass Tubing: Do not touch heated glass until it has time to cool. Hot
glass looks just like cool glass. To remove stoppers from glass tubing or thermometers,
grasp tubing close to stopper and push gently with twisting. Use water or glycerin for
lubrication.


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5. Laboratory Dress: Pull hair back and wear eye protection when required. Sleeves that
are too tight prevent freedom of movement, whereas sleeves that are too loose may cause
you to overturn apparatus or glassware. Aprons protect clothing from corrosive or
staining chemicals. Gloves protect hands from corrosive chemicals. Handle hot objects
with insulated gloves. Do not wear open-toe shoes that allow spilled chemicals or broken
glass to come in contact with your feet.
6. Working with Test Tubes: Gently heat solids or liquids in a test tube near the liquid or
solid surface. Be prepared to remove the tube from heat quickly to prevent eruption.
Never point a test tube or reaction vessel at another person. For safety and neatness, place
test tubes in a rack.
7. Chemicals in the Eye: Rapid treatment is vital. Run large volumes of water over eyeball
until medical help is available. Wash with large volumes of water for at least 15 minutes.
Alkaline materials in the eye are extremely hazardous. Know the location of the
emergency eyewash station.
8. Safety Shower: Use this for chemical spills or a fire victim. Operate by pulling down on
ring and keep the area near the shower clear at all times. Remove clothing from area
affected by spills.
9. Fire on Clothing: Do not run or fan flames. Smother fire by wrapping victim in fire
blanket or lab coat and use the shower or a carbon dioxide fire extinguisher.
Extinguishing a Fire using a fire extinguisher:
(1) Know its location
(2) Remove from mounting
(3) Pull pin
(4) Squeeze lever
(5) Discharge at base of flame
(6) Report use and recharge
(7) Use dry sand to extinguish burning metals
10. Unauthorized Experiments: Always work under instructor's or lab technician's
supervision in the laboratory.


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11. Eye Protection: Normal eyeglasses are usually not adequate. Do not wear contact lenses
in the lab. Eye protection is especially important when working with corrosive materials
and vacuum and high pressure apparatus.
12. Acid/Alkali Spills: For acid spills, use solid sodium bicarbonate followed by water. For
alkali spills, wash with water followed by dilute acetic acid.
13. Handling Flammable Liquids: Flammable liquids should always be stored in an
approved storage cabinet. Extinguish all flames in the area where flammable solvents are
used, as vapors may travel to ignition source and flash back.
14. Handling Mercury: Mercury spills are very hazardous. Droplets should be picked up by
suction and a mercury spill kit used to complete cleanup. Notify lab technician
immediately when mercury spills occur.
15. Protection from Toxic Gases: Emergency air masks should be used. However, because
our lab is not equipped with such masks, clear the area where gases are, and notify the lab
technician.
16. Waste Disposal: Hot glassware or reactive chemicals should be discarded in a
nonmetallic container separate from paper and other flammable waste. Test-tube
quantities of hazardous liquids can be flushed down the sink with plenty of water.
Contact lab technician for disposal of large quantities of hazardous materials or anytime
you are not sure of how something should be disposed of.
17. Labeling Chemicals: All chemicals should be clearly labeled. Do not use materials from
unlabeled containers. Avoid contamination. Never return reagents to their container.
Clearly label chemicals as you work.
18. Carrying Chemicals and Equipment: Carry long apparatus such as tubing or burets, in
an upright position close to the body. Grasp bottles firmly with both hands and hold them
close to the body. Do not carry bottles by the neck. Use a bottle carrier when transporting
chemicals any distance.
19. Transferring Liquids: Remember, Acid to Water. Do not pipette by mouth, use a bulb.
Use gloves when pouring corrosive liquids. Use a funnel when filling a bottle or flask and
prevent an air block by raising the funnel. Pour hazardous liquids over a sink.
20. Fume Hood: Use a fume hood equipped with a safety glass when working with toxic or
flammable materials.


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21. Gas Cylinders: Protect cylinder valve with cap. Fasten cylinders securely. Transport
cylinders on a hand truck, don't roll. Do not drop cylinders. Mark cylinders when empty.
22. Handling Sodium and Potassium: Fire or explosion may result when metallic Na or K
are exposed to water. Store them under light oil. Metal can be cut safely with a spatula on
a paper towel. Destroy residues with alcohol. Cool if necessary.
Prohibited actions in the lab:
1. No smoking
2. No foods or beverages
3. Unauthorized touching and misplacing of equipments, reagents, and chemicals
4. Unauthorized operation of any equipment in the lab is prohibited
5. Don’t leave any experiment unattended
6. Unnecessary movement in the lab is forbidden
Pay attention for any activity in the lab and always be prepared to help fellow students
in an emergency
I read the safety rules in this lab and agree to work in the lab accordingly.
Name……………………………………signature…………………..date………………..


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PRACTICAL 2: Sample collection and preparation for chemical analyses
Modern methods of chemical analysis require small amount of that must be collected and
prepared in such a way that gives the best reasonable estimate of the total batch. This is
because it is for example not practically feasible to analyze the whole animal body, even
of small animal. If we are interested in the protein content of hay produced from a field,
we certainly can’t grind all the hay produced, even one bale would tax the facility of most
laboratories. We therefore resort to the use of core sample taken from as many different
bales as is reasonable.
Small quantities of sample to be analyzed to be taken from as many part of the pile as are
accessible so that the proportion of course and fine particles in each small amount will be
the same as in the total amount of feed being sampled.
♣ ote!!
In general Samples must be taken at random. It must be thoroughly mixed. Then
divided them in to quarters, eights, sixteen’s etc. as desired.
A. Obtain a small quantity from several location within a lot of feed
B. Finely grind the sample to pass l mm sieve size and mix – until each spoonful is
representative of every other spoonful within the sample as whole as of the overall lot of
feed.
C. Keep the sample in a tightly closed container and label it
Sample Type, Species, Part of Plant/Animal, Date of Collection, Place of Collection
(Town, Altitude, Latitude, Rain Fall etc.)
♣ Sampling Areas
1. Sampling from field: - for pasture/grazed forage
Ö Using quadrant (of different size). The pasture is cut from as many site as possible


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2. Sampling from the storage
In this case mostly dried feeds such as cereal grains, Silage, Hay etc.
- Use long pipe to take samples
- Take randomly from different length and side of standing sack.
- Mix it thoroughly
- Pile it up and make flat at the top
- Quarter it and discard two diagonally opposite quarters
- Apply quartering up to the maximum limit of minimization
3. Liquid (Urine, Blood)
Liquids are more homogeneous than solids.
Record Incoming Samples
Upon receipt at the laboratory samples are assigned batch numbers. The requester has to
fill a form providing sample description, source, requested analyses, costs for each
analysis and budget code for charges. This form must be approval by the lab manager for
charge back payment. The Animal Nutrition Laboratory uses this information to build up
a data base on results of analyses and source description of all samples that pass through
the laboratory.
♣ Grinding Feed Samples
Homogeneous powders from dried samples are required for most chemical analysis. The
most frequently used grinder is hummer mill; brush, sieves and trays are accessories. It is
important to thoroughly clean the mill and the bags after each grinding to avoid cross
contamination. The particle size of the ground material can be important but for general


17
routine analysis, samples milled to pass through a 1mm mesh sieves are usually
satisfactory. For certain determination, e.g. Nylon Bag degradability, the sample should
be ground to pass through a 2mm mesh sieve. The ground sample is stored in plastic
containers with lids to keep out dust.
♣ Preservation (Wet Sample)
Freeze drying (Sample for polyphenolics determination)
Deep Freezer (Fecal Samples)
Preservation in acid media (Rumen Fluid and Urine)1
10 drops of H2SO4 in 100 ml, Store in cold room 4o
C in airtight plastic container.
Sample for VFA analysis: Immediate analysis for freezing


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Practical 3: Writing laboratory reports as scientific paper
Writing a scientific paper is one of the main duties of an animal scientist. They must not
only perform research experiments to test new ideas that will hopefully improve animal
production and meat quality; they have to publish these results as well. This section of
this laboratory is to provide students with a guide to writing a scientific paper. Within
this section you will find key components that are necessary to include in scientific
papers. We will also discuss helpful hints that may make your writing easier.
Procedures of scientific laboratory reports
1. Title
2. Objectives of the experiment
3. Introduction
4. Materials and Methods
5. Procedure
6. Results
Tables/charts/graph format
7. Statistical analysis of data
8. JAS format for tables and graphs
Describe the data in text format as well
9. Discussion
Interpret your results
Relevance to hypothesis?
Significance?
Compare to previous research if available


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10. Conclusion
Summarize your findings
How does it correspond to the original objectives of the lab
Were they met?
Why or why not?
Implications?
Attach a copy of data collection sheets
11. Recommendation


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Practical 4: Determination of Partial Dry matter and moisture content
Principle:
The moisture content in the foodstuff is determined by drying the sample in an oven at
1050
C overnight or 135 0
C for 2 hr - until constant weight loss in weight of the sample
gives amount of moisture.
This method is satisfactory for most feedstuffs but with a few, such as silage, significant
losses of volatile material may take place in oven drying. Such substance will obviously
be counted as water.
For materials containing much sugar or hemicelluloses, the temperature should not
exceed 700
c and drying is preferably conducted in a vacuum oven under reduced pressure
The DM determined from these type of drying is Partial DM.
Apparatus
Drying Oven
Metal plate/ Paper bag
Weighing balance
Procedure
- Weight your metal plate/ Paper bag and label it.
- Weight out a small quantity of the prepared sample in to pre-weighed container.
- Dry in an appropriate over until there is no further loss in weight.
- Weigh the sample after drying.
- Calculate the percentage of moisture or dry matter as follows.
% Moisture =

% Partial DM =
% Partial DM= 100% - % moisture
Species
Plant/Animal
Part of
plant/Animal
Date of
collection
Wt of Empty
Container(gm)
Wt of fresh
Sample(gm)
%
Moisture
%
Partial
DM
Review Question
1. The moisture content the forage supplied to a steer is 40%. If the dry matter requirement
of the steer is 6 kg/day. How many kg of the forage should be given to the animal in
order to satisfy its DM requirement?
2. What is the drawback of high- Temperature drying to determine DM of a green feed?
3. Assume that an air dry sample of wheat weighs 74gm.after the sample was heated for
several hours in an oven at 1000
C the weight changed to 65gm. Calculate the percent
water and percent DM content of the sample.


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Practical 5: Determination of absolute dry matter and ash
Principle: the dry matter content of feed sample and other material is expressed on three
dry matter basis: as fed, partially dry and dry.
As fed: refers to the feed as it is consumed by the animal, the term as collected is used for
material which are not usually fed to the animal, i.e urine, faces etc. if the analysis on a
sample is affected by partial drying, the analyses are made on the as fed or as collected
sample.
Partially dry: reports to a sample of as fed or as collected material that has been dried
in an oven at a temperature usually about 600
C or freeze-dried and has been equilibrated
with the air. This analysis is referred to as partial dry matter percent of as fed or as
collected sample. The partial dry sample must be analyzed for dry matter (determined in
an oven at 1050
C) to correct subsequent chemical analyses of the samples to a dry
basis.
Oven dry: refers to a sample of material that has been dried at 1050
C until all the
moisture has been removed. Similar term 100 per cent dry matter or moisture free is also
used. if dry matter (in an oven at 1050
C) is determined on an as fed sample, it is
referred to as dry matter of as feed sample. if dry matter is determine on a partial dry
sample it is referred to as dry matter of partially dry sample. it is recommended that
analyses be reported on the oven dry basis (100 per cent or moisture free).


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Sample as fed or as collected
↓ ↓
Sample contain more than 88% dry matter sample contain less than 88%
dry matter
↓ ↓
Grind, using a 1mm sieve Determine partial dry matter %
on as fed sample
↓ ↓
Determine dry matter directly at 1050
C dry in an oven at 600
C or freeze-
dry
↓ ↓
This is known as: as fed dry matter This analysis is referred to as
partial dry matter of
As fed or as collected sample
↓
Grind immediately, using 1mm sieve
↓
Determine dry matter at 1050
C. This analysis is
referred to as dry matter % of partial dry
sample
As fed Air dry Oven dry
%Water May be any % Usually 10% 0%
%Dry Matter (%CP, %
EE, %CF, %NFE, %Ash)
100% - % water Usually 90% 100%


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Composition figure expressed on the basis may be converted to another basis by the use a
sample ratio, as follows:
=
Apparatus:
- Drying oven
- Crucible
- Weighing scale (sensitive to 0.01gm)
- Desiccators
Procedure
- Hot weigh an empty crucible (W1) and transfer 0.5 to 1gm (W2) of partially dried (air
dried) sample in to it.
- Place the crucible with the sample in the oven at 1050
C overnight
- Weigh the crucible with the dried sample (W3)
% Absolute DM = Where: W1 - weight of empty crucible
W2 - weight of original sample
W3 - weight of dried sample + crucible
Wt of
Crucible
(gm)
Wt of air
dried sample
(gm)
Wt of oven
dried sample
+Crucible (gm)
%
DM
Wt Ash+
Crucible
(gm)
% Ash %OM


25
Ash determination
The ash content is determined by ignition of a known weight of the sample at 5500
C-
6000
C until all carbon has been removed for 3-4 hrs. The residue is the ash and is taken to
represent the inorganic constituent of the sample.
Apparatus:
- Muffle Furnace
- Porcelain crucible or 50ml Pyrex beaker
- Weighing scale (sensitive to 0.01gm)
- Desiccators
Procedure:
- Hot weigh an empty crucible (W1) and transfer 0.5 - 1gm (W2) sample in to it. The
material left after DM determination could also be used as a sample.
- Place in a muffle furnace and ash the material at 5000
C overnight
- Hot weigh the crucible with ash (W3)
% Ash = Where: W1 - weight of empty crucible
W2 - weight of original sample
W3 - weight of ash + crucible
Review Question
1. If the feed contains 80%DM on air-dry basis and 3% NFE, what would the percentage of
NFE on an oven-dry basis?
2. Discuss the three dry matter bases in which the composition of feed may be expressed.
3. Describe about Proximate (Weende) feed analysis and Detergent (Van Soest) System of
feed Analysis.


26
Practical 6: Determination of crude fat (Ether extract)
Principle:
Estimation of EE in feeds may be made either by weighing the extracted material after
evaporating the solvent or by recording the loss in wt of the moisture free sample
following its extraction by anhydrous ether.
The crude fat is determined by subjecting the sample to a continuous extraction with
petroleum ether for a defined period. The residue after evaporation of the organic solvent
is the crude fat or Ether Extract (EE)
While the crude fat in most feeds is usually mostly true fats, it may also include varying
amount of other ether-soluble internals such as waxes, organic acids, sterols,
phospholipids, fat-soluble vitamins, carotene, chlorophyll pigment etc. hence, the
designation “crude” fat. The amounts of ether- soluble materials in a feed, which are not
true fats, however, usually represent only a very small percentage of the overall feed.
Reagents:
Chloroform/Benzene/Petroleum ether (bp. 400
to 600
C)
Apparatus:
Soxhlet Extractor
Boiling flask with ground joint
Extraction thimble
Hot plate
Weighing balance
Oven
Filter paper


27
Procedure
1. Weigh out a small quantity of the prepared (previously dried) sample (usually less than
5g) on a pre-weighed filter paper and wrap it, tie it with thread so the filter paper may not
be opened easily during extraction.
2. Take the clean, dry boiling flask and weight it accurately
3. Put the wrapped sample in an extraction thimble and plug it with fat free cotton.
4. Place the thimble with the sample into the soxhlet and fill soxhlet with organic solvent by
pouring it through the condenser at the top by means of a glass funnel. (the amount of
solvent taken is about 11
/2 times the capacity of the extractor)
5. Extract for 3 to 5 hours. After extraction is over, remove the thimble with the sample
from soxhlet. Assemble the apparatus again heat it on the hot plate to recover all the
organic solvent from the boiling flask.
6. Put the boiling flask in a hot air oven 1000
C for 1 hour, cool in a desiccator and weigh (to
remove water).
7. Calculate the amount of crude fat by difference.
% Crude fat
x Preserve the extraction thimble with the material in a desiccator for CF determination.
Wt of empty
Receiver (gm)
Wt of air dried
sample (gm)
Wt of EE +
Crucible (gm)
% EE On DM
basis
Remark


28
Review Question
1. What is the justification for a crude fat (EE) to be a true fat?
2. What are the factors, which affect the chemical composition of feed?
3. What are the limitations associated with proximate analysis of feeds?


29
Practical 7: Determination of crude fiber content
Principle:
CF content is determined based on acid stomach digestion and subsequent alkaline
intestinal digestion of consumed food. CF is determined by subjecting the residual food
from EE to successive reaction boiling acid alkali of defined concentration.
Boiling the sample by dilute H2SO4 (1. 25%) and dilute NaOH (1.25%) to remove
protein, sugars, starches and more soluble hemicelluloses and mineral.
This fraction was designed to include those materials is a feed which are of low
digestibility. Included here are cellulose, certain hemicelluloses and some of lignin if
present.
Apparatus:
Filter paper, Buchner funnel, furnace, crucible, oven and vacuum pump, Beaker.
Reagents
1.25% H2SO4
1.25% NaOH
Ethyl alcohol
Procedure:
1. Weigh out the residue left after EE (practical 4) or prepare a fat free Sample and boil in
1.25% H2SO4 for 30 min and filter by suction through a Buchner funnel.
2. Wash the insoluble matter with boiling water.
3. Boil again with 1.25% NaOH for 30 min and filter by suction through Buchner funnel
and wash with boiling water until the washing is neutral to litmus paper.


30
4. Put the residue in crucible and dry it in oven (the residue consists of the crude fiber and
the more insoluble mineral matter of the feed sample)
5. After you dry it in an oven for one hr then weigh it.
6. Ash the residue to oxidize off the CF and weigh the ash.
7. Calculate the amount of crude fiber in the sample by subtracting the weight of the ash
from the wt of the residue in step 4.
8. Calculate the percentage of CF as follows
% CF
Wt of empty
Crucible (gm)
Wt of CF +
Crucible (gm)
Wt of Ash +
Crucible (gm)
% CF on
DM basis
Remark
Review Question
1. What do you understand if the sample contains high fiber content and vise versa?
2. What is the function of fiber with regard to digestion?


31
Practical 8: Determination of protein content (Total Nitrogen)
Principle:
Total nitrogen is determined using the kjeldahl method. Organic nitrogen is converted in
to ammonia ions by digestion with concentrated sulphuric acid in the presence of a
catalyst such as mixture of potassium (or sodium) sulphate with selenium or copper
sulphate. Following kjeldahl digestion, the digests made alkaline and Ammonium (NH4)
is determined either by steam distillation of Ammonia (NH3), which involves trapping in
boric acid, or by colorimetric (NH4) using an auto analyzer.
From this titration the amount of titration of nitrogen is determined; it is multiplied by
6.25 to convert to Crude Protein (CP). This conversion factor, found by determining the
amino acid content, is an average value for conversion of nitrogen for protein in most
plants. For specific grains, seeds, etc, other conversion factors have been determined.
2NH3 + H2SO4 (NH4)2SO4
OR
NH3 + HCl NH4Cl
Apparatus:
x Kjeldahl digestor
x Digestion tubes, 100ml
x Distillation rack
x Distilling unit Kjeltic system
x Mettler balance
x Automatic Burette


32
x Erlenmeyer flasks, 250ml
x Magnetic stirrer
Reagents:
x Conc. H2SO4 (98%), AR grade N-free
x Standard acid solution, 0.1N HCl 0.01M H2SO4
x Sodium hydroxide solution (35-40%)
x Methyl Red and Methyl Blue Indicator
x Boric acid (4%)
x Catalyst tablet: Selenium or mixture of Na2SO4 and CuSO4 in the ratio of 10:1 powdered
mixture can also be used.
Procedure
- Weigh 0.3g ground dried sample (if fresh faeces weigh 0.6g, if milk or urine take 1ml ) in
to a digestion tube
- Add 1gm catalyst mixture of K2SO4 and anhydrous CuSO4 in the ratio 10:1
- Add 5ml conc. H2SO4
- Place the digester and bring the temperature to 3500
C. Digest for about 1.5hours or until
the sample almost looks clear solution. If the digest is light brown or yellow, the
digestion may be incomplete requiring additional digestion time. If the digestion is
complete, the digest should be either clear or light blue.
- Remove the tube from the block digester and allow it to cool
- Carefully add 30ml of distilled water in to the digestion tube.
- Carefully add 40-50ml NaOH.
- The digestion mixture is now ready for ammonium determination by the steam
distillation procedure. The ammonium in digestion mixture can also be determined
calorimetrically using an auto-analyzer.
x Distillation and titration

Place the digestion tube in the tecator steam distillation apparatus. Insert the digestion
tube in the system and collect 150ml distillate in the receiver flask containing 25ml of 4%
boric acid.
Titrate the distillate against a standard acid (0.1N HCl). the colour change is from green
to steel- blue then pink with the end point occurring when one drop of acid result in a
colour change from steel-blue to light pink.
% N
%CP = % × F
Where VS = Volume of acid (HCl/H2SO4) consumed for sample titration
VB = Volume of acid (HCl/H2SO4) consumed for blank titration
N = Normality of HCl (H2SO4)
W = Weight of sample taken in gram
DM% = Dry matter percentage of the sample
F = conversion factor which is specific for different products/feeds
For example
F = 6.25 for all forages
F = 5.70 for wheat grains
F = 6.38 for milk
Review Question
1. Show how the factor of 6.25, which is used to estimate the percent crude Protein in a
feed, was obtained?

2. Suppose that a sample of oats grain has been analyzed by the proximate analysis method
and produce these results: 11% water, 3.2% ash, 405% EE, 11%CF, and 1.9%nitrogen.
Calculate the percentage of NFE.
Practical 9: The detergent system of Fiber analysis (Van Soest method)
Introduction
The fact that crude fiber/NFE system does not provide an accurate picture of the
carbohydrate fraction of feedstuffs, primarily because of solubilization of variable
amounts of hemicellulose and lignin in the crude fiber analysis, the detergent system of
feed analysis was developed by P. J. Van Soest and associates, working at the USDA
station at Beltsville, MD, a rapid technique of separating feed carbohydrates on the basis
of nutritional availability to ruminants and ruminal bacteria. Essentially, the method
divides feeds into two fractions: (1) plant cell contents, a highly digestible fraction
consisting of sugars, starches, soluble protein, pectin, and lipids; (2) plant cell wall
constituents, a fraction of variable digestibility consisting of insoluble protein,
hemicellulose, cellulose, lignin, and bound nitrogen. The method involves boiling a
sample in a neutral detergent solution. The soluble fraction is termed neutral detergent
solubles (cell contents), whereas the fibrous residue is called neutral detergent fiber (cell
wall constituents). Unlike crude fiber and NFE, both NDS and NDF accurately predict
the proportions of more and less digestible fractions, respectively, found in a wide variety
of feedstuffs.
The Van Soest (or detergent) scheme has been further refined with the addition of acid
detergent fiber (ADF) analysis, which breaks down NDF into a soluble fraction
containing primarily hemicellulose and some insoluble protein and an insoluble fraction
containing cellulose, lignin, and bound nitrogen. Furthermore, the content of lignin in
ADF can be determined by either treating the fiber with H2SO4 to dissolve the cellulose
or by oxidation with permanganate to degrade the lignin. This analysis is quite important
because lignin has been shown to be a major factor influencing the digestibility of
forages.


35
Besides accurately describing the chemistry of the feed, the results of this scheme are
highly related to digestibility and intake of feedstuffs. Among the chemical entities, the
Neutral detergent fiber (NDF) is highly related to intake and together with other entities
(Acid detergent fiber, lignin and silica). Prediction equations (summative) were
developed to estimate digestibility and energy value of feedstuffs. Except cost and
robustness, the detergent method is technical sound for feed resource characterization.
1. eutral Detergent Fiber (DF) Procedure
Reagents
x NDF solution: To 1 L of H2O add 30 g of sodium lauryl sulfate, 18.61 g of disodium
dihydrogen ethylene diamine tetraacetic dehydrate, 6.81 g of sodium borate
decahydrate, 4.56 g of disodium hydrogen phosphate and 10 mL of triethylene glycol.
x Amylase solution - Heat-stable α-amylase (Sigma No. A3306 – from the Dietary Fiber
Kit).
x Acetone
Materials
x Refluxing apparatus
x Whatman #541 filter paper
x Aluminum pans
Procedure
1. Place 0.5 to 1.0 g sample in 600-mL Berzelius beaker.
2. Add 100 mL of neutral detergent fiber solution.
3. Heat to boiling (5 to 10 min). Decrease heat as boiling begins. Boil for 60 min.
4. After 60 min, filter contents onto preweighed Whatman #541 filter paper under
vacuum. Use low vacuum at first, increasing only as more force is needed.
5. Rinse contents with hot water, filter, and repeat twice.
6. Wash twice with acetone.
7. Fold and place in pre-weighed aluminum pan.
8. Dry overnight in 100°C oven.

9. Cool in desiccator.
10. Weigh and record the weight.
ote. For samples with high starch content: Add 50 μL of heat-stable amylase to the
beaker along with NDF solution as in Step 2, and follow remaining steps. For the most
difficult samples, a 1-g sample can first be treated with 30 mL of 8 M urea solution plus
50 Μl of heat-stable amylase solution. The mixture can be heated on a steam bath at 80 to
900C for 5 min, then incubated at room temperature for 4 h or overnight. After
incubation, add 100 mL of NDF solution and treat as in Step 3 and following. An
additional 50 μL of heat-stable amylase can be added at this point if desired.
2. Acid Detergent Fiber (ADF) Procedure
Materials:
x 600 mL Berzelius beakers
x Fiber digestion apparatus
x Sintered glass crucibles, 40 to 50 mL - coarse porosity
x Filtering flasks
Reagents:
x Acid Detergent Solution: Add 27.84 mL of conc. H2SO4 to 500 mL of distilled water
in a 1 L volumetric flask and bring to 1 L volume with water. Then and 20 g of
CTAB to the 1 L of acid solution.
x Acetone
Procedure
1. Transfer 1 g air-dried sample to Berzelius beaker.
2. Add 100 mL acid detergent solution.
3. Heat to boil (5 to 10 min), and boil exactly 60 min.


37
4. Filter with light suction into previously tared crucibles. Refer to procedure for matting
crucibles. #541 filter paper may be used instead of crucible if ADL is not being done.
You must pre-weigh filter paper first.
5. Wash with hot water 2 to 3 times.
6. Wash thoroughly with acetone until no further color is removed. Suction dry.
7. Dry in drying oven at 100°C for 24 h.
8. Cool in desiccators. Weigh and record weight.
3. Acid Detergent Lignin Procedure
Reagents:
x 72% H2SO4 standardized to specific gravity of 1.634 at 20°C.
Procedure
1. Place ADF crucible in a 50 mL beaker on a tray.
2. Cover contents of crucible with 72% H2SO4. (Fill approximately half way with acid.)
3. Stir contents with a glass rod to a smooth paste.
4. Leave rod in crucible, refill hourly for 3 h, stirring the contents of the crucible every
hour.
5. After 3 h, filter contents of crucible using low vacuum at first, increasing it only as
more force is needed.
6. Wash contents with hot water until free of acid (minimum of five times).
7. Rinse rod and remove.
8. Dry crucible in oven at 100°C for 24 h.
9. Cool in dessicator. Weigh and record weight.
10. Ash in muffle at 500°C for 4 h.


38
Practical 10: Determination of urea
The urea content of feed ingredients can be quantified with this technique.
Reagents
x Active charcoal.
x Carrez solution I: dissolve 21.9 g dehydrated zinc acetate in water; add 3 ml glacial acetic
acid and dilute to 100 ml with distilled water.
x Carrez solution II: dissolve 10.6 g potassium ferrocyanide in 100 ml distilled water.
x Chlorhydric acid 0.02N.
x Sodium acetate solution: dissolve 136g sodium acetate trihydrate in 1000 ml water.
x Solution of 4-di-methylaminobenzaldehyde (DMAB): dissolve 1.6 g DMAB in 100 ml
ethanol at 96% and add 10 ml chlorhydric acid (d = 1.18 g/ml).
x Standard urea solution: dissolve 0.1 g urea in 100 ml of water.
Material
x Rotary agitator
x Spectrophotometer with 10 mm cells.
Method
Weigh approximately 2 g of sample to within 0.001 g or an amount expected to contain
50–200 mg urea and place in a 500 ml volumetric flask. Add 150 ml chlorhydric acid
0.02N, agitate for 30 minutes then stir in 10 ml of the sodium acetate solution. Add 1g
active charcoal, shake thoroughly and let the mixture rest for 15 minutes. Add 5 ml
Carrez solution I, followed by 5 ml Carrez solution II, stirring well between the two


39
additions. Calibrate with distilled water and mix well. Filter one part through a dry filter
paper into a clean, dry 250 ml beaker.
Determination
Transfer 10 ml of filtrate to a test-tube with a ground glass stopper. Add 10 ml 4-DMAB
solution. Stir; rest for 15 minutes. Measure the absorption of the solution as indicated
above against a reference solution prepared with 10 ml 4-DMAB solution and 10 ml
water. Draw a graph relating absorption to the amount of urea present.
Expression of results
Determine the quantity of urea in the sample referring to the calibration curve drawn.
Express results as a percentage of the sample: % urea × 0.4665 = % N urea
NOTE:
If the sample is highly coloured, the amount of active charcoal should be increased to
above 5 g. The final solution after filtering should be colourless.


40
Practical 11: Determination of Uric acid
This method can determine uric acid content and its salts in both chicken manure and in
feed and ingredients.
Reagents
x Sodium hydroxide solution. Dissolve 50 g NaOH in 50 ml water, stir well and store
solution in a plastic container.
x Neutral formaldehyde solution:
Check the concentration of the solution available by mixing 30 ml of it with 50 ml NaOH
IN solution and 25 ml hydrogen peroxide solution (20 volumes). Heat in water-bath until
effervescence stops. Cool and titrate with NaCI IN using phenophthalein indicator.
Perform a blank titration using 3 ml water instead of formaldehyde and calculate the
concentration as follows:
1ml NaOH 1N solution = 0.03 g formaldehyde.
Where B = blank titration


41
T = titration of sample
Prepare a neutral solution with 17.5 g of formaldehyde, 250 ml water and 500 ml ethanol.
Adjust the pH to 7.0 with a solution of NaOH 0.1N. Dilute to 1,000 ml with water, stir
and adjust pH again if necessary.
x Buffer solution of succinate. Heating, dissolve 29.5 g succinic acid in 750 ml water and
20 ml of the NaOH solution. Let cool and add a sufficient amount of formol solution
containing 17.5 g formaldehyde, stir well and adjust pH to 6.0 with NaOH solution.
Dilute to 1,000 ml with distilled water, stir and adjust pH if necessary.
x Sodium thiosulphate solution. Dissolve 25g Na2S2O3 • 5H2O in 1,000 ml distilled water.
x Actol solution. Heating, dissolve 3 g actol in 50 ml distilled water and 1 ml lactic acid.
Dilute to 100 ml with water, filter and store in a dark bottle. Do not expose to direct light.
x Ammoniated magnesium solution. Dissolve 8.75 g of MgSO4 • 7H2O and 17.5 g NH4 Cl
in 50 ml water. Add 30 ml NH4OH (d = 0.88 g/ml), stir well and dilute to 100 ml with
water.
x Benedict and Hitchcock's reagent. Mix 35 ml actol solution with 15 ml ammoniated
magnesium solution. Add 50 ml ammonia hydroxide (d = 0.88 g/ml). Prepare
immediately before using.
x Standard uric acid solution. Weigh 250 mg uric acid to within 0.1 mg and transfer to a
round-bottomed flask coupled to a reflux condenser. Add 100 ml of the ethyl
formaldehyde solution and heat by reflux in a water-bath for 30 min, shaking frequently.
Cool and transfer to a 250 ml volumetric flask, washing the round flask with ethyl
formaldehyde sol. calibrate with this solution and mix. 1 ml contains 1 mg uric acid.
x Petroleum ether, boiling point 40–60°C.
Material and equipment
x Spectrophotometer with 10 mm quartz cells.
x Glass percolation tubes approximately 240 mm long, 18 mm inside diameter and
approximately 120 mm long at the bottom and 8 mm inside diameter.
Extraction of uric acid:


42
i. From chicken manure: weigh some 0.4 g dry manure to within 0.001 g and place in a 150
ml round-bottomed flask. Add 60 ml neutral formaldehyde solution, connect to a reflux
condenser and heat in water-bath for one hour. Cool and filter through a crucible
(porosity 4) into a 100 ml volumetric flask. Rinse the flask 3 times consecutively with 10
ml portions of the ethyl formaldehyde solution, filtering each quantity through the
crucible into the volumetric flask. Calibrate with the same solution and shake.
ii. From feed: weigh 4 to 5 g of sample to within 0.001 g and extract fat with petroleum
ether. Transfer the defatted sample quantitatively to a round-bottomed flask and remove
the remaining solvent with a gentle air current. Continue the analysis as above, beginning
with the addition of 60 ml ethyl formaldehyde solution.
Determination
Using a pipette, transfer 20 ml preweighed extract of sample according to the methods
described, to a 50 ml centrifuge tube. Add 10 ml Benedict and Hitchcock's reagent, mix
well and leave in dark to rest one hour. Centrifuge at 2,000 rpm for 15 min. Remove
supernatant and leave to drain for 10 min. carefully clean away any remaining liquid
without disturbing the precipitate and add 20 ml sodium thiosulphate. Dissolve the
precipitate, stirring with a thin glass rod. Using a pipette transfer 5 ml of this solution to a
200 ml volumetric flask containing 40ml succinate buffer solution, calibrate with distilled
water and mix well. Measure the absorption of the solution at 294 nm in 10 mm silica
cells compared against a solution prepared by mixing 5 ml sodium thiosulphate solution
with 40 ml succinate buffer solution brought to 200 ml with distilled water. Determine
the amount of uric acid by means of a calibration curve.
Calibration curve
Using a pipette, transfer 2, 4, 6, 8, 10 and 12 ml standard uric acid solution (equivalent to
similar amounts of uric acid in mg) into a series of 50 ml centrifuge tubes and bring them
up to 20 ml with the ethyl formaldehyde solution. Add 10 ml Benedict and Hitchcock's
reagent to each tube, stir well and leave in darkness for one hour. Continue the method as
for determination, from the centrifuge stage. Measure the absorption of the solution and


43
draw the calibration curve, plotting absorption (y) against the corresponding amount of
uric acid in mg (x).
Expression of results
The N content of the uric acid as a % of the sample is given by the formula:
WhereA = mg of uric acid (in the aliquot of extract of sample) determined by
photometric measurement.
W = weight of sample in g.


44
Practical 12: In vitro Digestibility
Principle
In vitro digestibility techniques provide a quick, inexpensive, and precise prediction of in
vivo or conventionally determined digestibility in ruminants. Essentially, the method
simulates the processes that occur in the rumen, and probably the most commonly used in
vitro technique is the one devised by Tilley and Terry (1963). Although the original
Tilley and Terry procedure has been modified by many researchers, the basic procedure
is the same as that used in the Animal Nutrition Laboratory. A copy of the procedure is
provided with this chapter. Estimates of digestibility by the Tilley and Terry procedure
are within 1 to 3% of conventionally determined values. The in vitro procedure does a
better job of prediction than chemical composition because it accounts for all factors
affecting digestibility, whether known or unknown, which is not possible with current
chemical methods. This accounting for additional factors is primarily a function of the
use of ruminal fluid from a donor steer as the digestive agent, thereby including unknown
factors that simple chemical analyses of the feed do not reveal. As indicated previously,
the in vitro procedure is quite simple, but nonetheless subject to a number of variables
that may influence the results obtained. Basically, a small sample of feed (~.5 g) is
weighed into a 50-mL centrifuge tube. McDougall's buffer (based on the composition
of sheep saliva) and ruminal fluid from a donor animal are added, and the tube is allowed
to incubate for 48 h at 390C. The fermentation is then stopped, tubes are centrifuged, and
supernatant fluid discarded. Acidified pepsin is added, and the tube is allowed to incubate


45
for another 48 h at 390C. Finally, the contents are filtered, and the residue is dried and
weighed. In vitro dry matter disappearance is determined by the following formula:
IVDMD % = 100 x [(initial dry sample wt - (residue - blank))/initial dry sample wt]
The blank value is determined by incubating a tube containing ruminal fluid and buffer,
without any feed sample. This accounts for indigestible materials introduced into the
vessel by the ruminal fluid inoculum that should not be counted against the feed.
The procedure is useful in that estimates of digestibility can be obtained in a few days on
a large number of samples. However, we should consider some of the variables that
influence the procedure. Four major ones are listed below from Johnson (1969).
A. Variations in the microbial population
1. Diet of donor animal
2. Animal to animal differences
3. Inoculum processing
B. Variations resulting from different storage, grinding, and processing
techniques in sample preparation
C. Differences attributable to the fermentation medium
1. Sample: inoculum ratio
2. Buffer
3. Nutrients in medium
D. Procedural variations such as length of fermentation and laboratory errors.
Given these variations, one can develop methods to standardize the in vitro
procedure. The largest source of variation among the four major sources listed above
is the variation in the microbial population. Difficulties associated with this source of
variation can be partially overcome by using more than one animal and by feeding
donor animals the same or a similar diet to that being evaluated in the in vitro system.
In addition, fluid should be removed at a standard time after feeding. The best


46
preference is to remove fluid ~4 h after feeding when microbial numbers are
maximal; however, several researchers prefer removing fluid after the donor has been
withheld from feed and water for 12 to 14 h. Generally, the standard method of
processing the inoculum is simply to strain whole ruminal contents through at least
four layers of cheesecloth. With regard to variations in the sample, it has been
observed that finely ground samples are more highly digested than are coarsely
ground samples. Hence, all samples should be ground in the same manner, and
grinding to pass a 1- to 2-mm screen usually is adequate. A commonly used sample:
inoculum: buffer ratio is 0.5 g:8 mL of ruminal fluid:32 mL McDougall's buffer.
Some laboratories add nutrients to the buffer, the most common being urea, to
prevent nutrient deficiencies during fermentation. Fermentation should be maintained
at a pH of 6.9 to 7.1 for optimum results. Procedural variations can be minimized by
standardizing temperature, time of fermentation, centrifugation speeds, and so on. In
fact, the key to successful in vitro analysis is to standardize as much as possible. In
this regard, it is a good practice to include a standard forage with each in vitro run as
a means of determining the validity of individual runs. All things considered; the in
vitro digestibility technique is the best means of laboratory evaluation of digestibility
available today. The procedure will continue to be used extensively for some time to
come. Students are referred to Johnson (1969) for further reading on the in vitro
technique.
IN VITRO PROCEDURE
Reagents
x McDougall's artificial saliva (mix four parts McDougall's to one part ruminal
fluid.)
x 9.8 g NaHCO3/L
x 7.0 g Na2HPO·7H2O/L or use 3.71 g anhydrous/liter
x 0.57 g KCl/L
x 0.47 g NaCl/L
x 0.12 g MgSO4·7H2O/L


47
x 4% (wt/vol) CaCl2 solution: 4 g CaCl2/100 mL
Mix the first five chemicals in 500 mL of water and stir until dissolved. Add remainder of
water. Before using, add in the 4% CaCl2 solution (use 1 mL of the 4% CaCl2 solution
per 1 L). Place the McDougall's solution, after the addition of the 4% CaCl2 solution,
into the 390C water bath and bubble in CO2 gas until the pH of the McDougall's solution
reads 6.8 to 7.0. When using the CO2 tank, open the top release valve, and then open the
smaller valve to release CO2 into the plastic line. After you have finished, close the top
release valve and close the smaller line release valve. Failure to close both valves result
in emptying the tank. When the tank reads at 50 lb of pressure, please tell the technician
so a new CO2 tank can be ordered.
Pepsin solution
x 6.6 g of 1:3,000 pepsin
x 100 mL of 1 N HCl (to prepare 1 N HCl add 80.4 mL of HCl/L of H2O)
Add deionized H2O to 1 L
PROCEDURE
1. Weigh out a 0.5-g sample and place into a labeled 50-mL centrifuge tube.
2. To this tube, add 28 mL of the McDougall's solution. Prewarm McDougall's in 390C H2O
bath. Add 7 mL of ruminal fluid (can alter quantity, but use 4:1 ratio of buffer to ruminal
fluid).
Place ruminal fluid on stir plate to avoid settling. Ruminal fluid is strained through four
layers cheesecloth before use. If possible, ruminal fluid should be obtained from at least
two animals.
3. Flush tube with CO2 (gently so sample is not blown out). Place cap on tube, invert several
times to suspend the sample, then place tubes into a rack, and place the rack into a 390C
water bath.


48
4. Also include at least four blanks (tubes containing no sample and 35 mL of the
McDougall’s to ruminal fluid mixture). Include two blanks per time interval if rates of
digestion are to be determined. Include 0.5-g samples of lab standards.
5. Incubate the tubes for 48 h.
6. Invert the tubes at 2, 4, 20, and 28 h after initiation of incubation to suspend the sample.
7. After 48 h of incubation, remove the tubes from the water bath. Centrifuge for 15 min at
2,000 x g and suction off the liquid by vacuum. At this point, one may freeze samples
until they can be filtered or until the pepsin digestion can be completed.
8. If you are doing the acid pepsin digestion, mix the pepsin solution, and add 35 mL of
pepsin solution to each tube. Incubate for 48 h in a 390C water bath, shaking at 2, 4, and 6
h after pepsin addition.
9. After the completion of the digestion (either McDougall's and ruminal fluid or the pepsin
solution digestion), filter your samples using the modified Buchner funnel and ashless
filter paper.
10. Dry the filter paper containing the sample in an aluminum pan for 12 to 24 h. Record
weights.
11. Ash each sample and record the weights. Ash at 500°C for 4 h.
12. Complete calculations.

Practical 13: The nylon bag technique (In sacco degradability)
Principle:
The nylon bay technique is used for measuring feed degradability in ruminants. In this
technique, a known weight of a sample is placed in bogs made of indigestible material.
The bogs are sealed tightly and placed in the rumen of fistulated animals and removed
after different periods of time (4 to 120 hrs), washed, dried and weighed. Degradability of
the substrate is determined by wt loss during the incubation periods.
This technique provides a means of ranking feeds according to the rate and extent of
degradation of DM, organic matter and nitrogen depending on the need.
Factors affecting Nylon bag degradation include:
- Placement of bogs in the rumen
- Particle size of the sample vis- a- vis the pore size of the nylon bay.
- Methods of washing.
- The length of time that the sample are incubated in the rumen
- The rumen environment in which degradability is determined (for the purpose of ranking
feeds according to their nutritive value, the diet of fistulated animal should be adequate in
rumen degradable nitrogen (N) and there has to be consistent feeding regimen)
General description
The total time for complete degradation varies with material being incubated and so d
other intermediate time chosen. As a rough guide concentrate requires 12-36 hours, good


50
quality forage 24-60 hours and poor-quality roughage 48-72 hours. These are times
recommended for reaching the potential degradation.
The pores in the bags not only allow the movement of the microbes and fermented
products, but also inert passage of fine particles. Passage of feed particles from the bags
without breakdown by rumen particles is corrected by using zero hour bags. These bags
are filled with the sample but are not incubated in the rumen. They are washed and dried
in the same way as the incubated bags. The zero hour bags are also used as a means of
correcting for passage of material from pressure applied to nylon bags during washing.
40-60 bags can be incubated in cattle while in sheep, 8-10 bags could be incubated at the
same time.
On removal from the rumen, the tied bags are healed by the necks and vigorously shaken
in a bucket of water. The string is cut (to detach the bag from PVC tubing) to clear
derbies trapped by folded material the bag and contents are rinsed in a washing machine
(individual bags remain tied). Alternatively, the bag material can be cleaned under
running tap water while rubbing between fingers and thumbs until the water run clear.
The bags are then dried to a constant weight at 60-700
C and the percent dry matter loss
calculated.
Materials:
- Nylon- bay (pore size 30-50µ and dimension 6.5 x 14cm)
- Nylon string or cord or plaster tube
- Analytical balance
- Drying oven
- Fistulated animal (sheep, cattle at least 3 at a time)
- Desiccators
Procedure:
1. Grind feeds thoroughly at 2mm screen (mesh size)
2. Determine the DM of the sample at 1050
C


51
3. Place about 2 gm sample in each nylon bay (which could vary with the type of
sample and total surface area of the nylon bag)
4. Tie the bags with about 50cm of nylon colder plastic tube and placed the material deep in
the rumen of frstulted animals
5. Incubate sample for 0,6,12.24,48, 72, 96, and 120 hrs and wash respectively using
running tap water or washing machine and dry nylon bays with residue at 60-700
C for
48hrs
6. Calculate DM disappearance
DM disappearance (%)
OR
DM disappearance (%)
Where Swa - weight of original sample + nylon bag
Swb - weight of sample + nylon bag after incubation
DMa - dry matter of feed sample
DMb - dry matter of residue sample
Bw - weight of empty nylon bag
REVIEW QUESTIO
A. Mention the different methods, which are used to determine digestibility.
B. Show how the factor of 6.25, which is used to estimate the percent crude Protein in a
feed, was obtained?
C. Suppose that a sample of oats grain has been analyzed by the proximate analysis method
and produce these results: 11% water, 3.2% ash, 405% EE, 11%CF, and 1.9%nitrogen.
Calculate the percentage of NFE.


52
Practical 14: Gas Production Test (Hohenheim Method)
Principle
The gas production test is based on the association between rumen fermentation and gas
production.
The in vitro gas production method can be used to measure the metabolizable energy of feeds
and to quantify utilization of nutrients.
Operation procedure
A. Substrate preparation
umber the gas syringes
Accurately weigh about 1g of substrate (maize straw and alfalfa hay), and transfer it
to the gas syringes
Preparation of Buffer medium


53
solution A˖
CaCl2.2H2O 13.2g;
MnCl2.4H2O 10.0g;
CoCl2.6H2O 1.0g;
FeCl3.6H2O 8.0g;
Add distilled water to 100ml
solution B˖
NH4HCO3 4.0g;
NaHCO3 35.0g;
solution C˖
Na2HPO4.12H2O 9.45g
KH2PO4 6.2g
MgSO4.7H2O 0.6g
solution D˖
100mg resazurin dissolved in
100ml distilled water
solution E˖
Cysteine HCl 625mg, Distilled water 95ml, 1M NaOH 4.0ml, Na2S
9.H2O 625mg
Order փ〟˄ml˅
1 distilled water 520.2
2 solution B 208.1
3 solution C 208.1
4 solution A 0.1
5 solution D 1.0
6 solution E 62.5
Ruminal fluid collection and inoculum
1. Mixed rumen content was squeezed through four layers of cheesecloth into a flask
under CO2 in a water bath kept at 39◦
C.
D. Procedure
x Weigh 200 mg of feed sample (1 mm ground) and insert carefully in 100 ml calibrated
glass syringe
x Include blank syringes without feed
x Include syringes with standard feed (concentrate or/and hay)
x Keep syringes overnight in circulating water bath at 39 °C

x Prepare fermentation buffer solution except rumen fluid and reducing solution in 2 l
bottle
x Keep bottle in water bath at 39 °C with stirrer overnight.
x Next morning
x Keep under CO2
x Add reducing solution and wait 20 minutes until color changes from blue to purple to
colorless
x Add rumen fluid
x Keep stirring under CO2 for 10 minutes
x Fill 30 ml into glass syringes
x Incubate syringes in 39 °C
x Shake syringes every hour during first four hours, then twice every hour
x Record gas production at 8 hours and push back piston to 30 ml if
gas production exceeds 70 ml

Operational flowchart


56


57
Measurement of Methane Production
Methane production after 24 and 72 hours of incubation can be measured using the
procedure described by Fievez, et al. (20)10. For measuring methane production at the
end of incubations, and after recording the final gas volume, the lower end of the
syringe will be connected to the lower end of another syringe containing 4.0 ml of
NaOH (10 M), which will be then introduced from the latter into the incubated contents,
thereby avoiding gas escape. Mixing of the contents with NaOH allowed absorption of
CO2, with the gas volume remaining in the syringe considered to be CH4. Net methane
and gas productions will be calculated by the differences of the methane and total gas in
the test syringe and the corresponding blank. The methane concentration will be
calculated as Jayanegara et al. (200):
Methane concentration = Net methane production/Net gas production.
Precaution: Sample of rumen liquor should be drawn after 2-3 h of feeding. Anaerobic
condition must be maintained throughout the experiment.


58
Practical 15: Conventional Digestion /metabolic experiments
Principles
The conventional digestion trial is the most reliable method of measuring the digestibility
of a feedstuff. Unfortunately, however, it is somewhat time consuming, tedious, and
costly. Basically, the feed in question is fed in known quantities to an animal. Usually,
the animal is restrained in an individual cage so that a quantitative collection of feces can
be made. Accurate records of feed intake, refusals and fecal output are kept, and a
subsample of each (usually 10% of daily output in the case of feces) is retained for
analysis. When estimates of nitrogen balance are desired, urine output also is measured.
Three animals per feed are required as a minimum, with more animals preferred. The
animals are usually allowed from 7 to 14 d to adjust to the feed and their cages, followed
by a 5- to 7-d period in which feces is quantitatively collected and feed intake is
recorded. Samples can then be dried, ground, and analyzed for the nutrients of interest.
Digestibility of any given nutrient can be calculated as follows:
utrient digestibility = [(nutrient intake - nutrient in feces)/nutrient intake] x 100
Even though conventional digestion trials are the standard with which all other measures
of digestibility are compared, the values obtained still vary ± 1 to 3 % as a result of
animal-to animal variation and sampling and analytical errors. Thus, it is clear why
nutritionists would seek alternative measures of digestibility because of the time and
expense of conventional trials.
A typical protocol for nutrient balance trials is as follows:
Procedures and General Consideration for utrient Balance Trials with Ruminants
I. Selection and care of animals
A. Select animals of similar size, sex, age, weight, and breeding. They should be free of
parasites, healthy, and vigorous; but not unduly excitable.


59
B. Place animals in preliminary feeding pens, and allow them to become adjusted to
environment and to the routines of handling and feeding. Gradually change the diets
to those that will be used in the trial. During this adjustment period, replace animals
that are poor eaters, and those that are overly excitable or otherwise unsuitable.
Weigh the animals at regular intervals, and keep a record of these preliminary
weights.
C. Allow the animals a few days for adjustment to metabolism cages after their transfer
from the preliminary holding pens. Protect them from flies and pests, keep them
clean, and provide for their comfort within the limits of the trial objectives.
D. Keep a daily record of temperature, weather conditions, unusual events in the animal
laboratory, and any observations of pertinence to animal behavior or performance.
II. Experimental Feeds
A. During the preliminary period, calculate the amount of diets (roughage, concentrates,
and supplements) necessary for the entire period of the study. Trials with ruminants
should consist of at least a 10-d adjustment period followed by a collection period
of 10 d. It is wise to provide a surplus of all dietary ingredients.
B. Ensure that all feed ingredients are uniformly mixed, so that ingredients fed initially
are comparable to those fed finally. In the case of concentrate mixtures and
supplements, mix amounts sufficient to provide at least 1¼ to 1½ times the amount
calculated to be fed during the actual trial. This allows for repeat days, spillage,
and so on.
C. Store all feed ingredients in covered containers conspicuously labeled and
completely labeled:
1. Project number
2. Trial number
3. Animal number (to receive diets)
4. Dates of trial
5. Experimenter


60
D. Secure the labels on containers! (ever label lids only). Situate feed cans
conveniently, and protect against inadvertent interchange of locations. For
roughages, select a quantity of feed that is in excess of needs, uniform in quality,
and properly identified. Store conveniently, and protect from contamination
specially dusts from feedstuffs and chemicals.
E. Preliminary sampling and analyses. Take a representative sample of all feedstuffs
and analyze before the onset of the actual trial to ensure that composition of diets
is as prescribed for the trial.
III. Detail Procedures
A. Check the accuracy and precision of all scales, balances, and volumetric equipment to
be used in measurement of feedstuffs and excreta.
B. Plan in detail a definite routine to be used in the trial - both for the preparation and
provision of diets and for the collection of excreta - and follow this routine
throughout. Generally, this involves weighing dietary ingredients directly into
individual feed pans, and, if necessary, mixing each animal's diet by hand to ensure
uniform distribution of ingredients and to prevent selective consumption by the
animal.
C. During the preliminary period, while animals are in metabolism cages, collect daily a
running grab sample from each of the dietary ingredients. Composite these into
single, 10-d samples for each of the diets. Label thoroughly! Use the same procedure
to collect additional 10-d, composite samples for each of the ingredients fed during
the subsequent 10-d collection period. LABEL COMPLETELY.
D. Feed consumed during the adjustment period will be excreted during the subsequent
collection period. Therefore, the daily feed must be kept constant throughout the 20-
d feeding period - constant in amount, and unvarying in composition.
E. Maintain a Barn Record showing the daily feeding performance of each animal, the
daily excreta, and any notes of consequence.
F. At the onset of the collection period, start the collection of urine and fecal samples
at a specified time, and strive to meet the same schedule throughout the trial. Be sure
to end the trial at the same time on the final day of collection.


61
G. If there should be any feed uneaten from feeding time to feeding time, remove the
entire refused portion, weigh and record the weight, and collect a representative
sample for analysis. LABEL COMPLETELY.
H. Collection of urine
1. Equipment used must afford quantitative collection of entire 24-h excreta. Collect
urine into clean containers that are safe-guarded against contamination from
feedstuffs, fecal material, and extraneous material (e.g., flies, bugs, dust, and so on).
Protect also from spillage and breakage (plastic jugs are preferred in most cases).
2. Add a few drops of toluene into the collection jugs at the onset of each 24-h period to
retard microbial activity (this will have insignificant effects on most analyses that are
conducted in routine trials).
3. When making the daily collections, provide a temporary container to collect any
excretion made during the process (a stitch in time…).
4. During the collection, suspend sediment by shaking the collection jug, and pour
quickly into a volumetric cylinder (graduated), and record the daily urine volume to ± l
mL. Test the reaction of the urine to litmus (note if basic) and determine specific
gravity and/or total solids by refractometry, if applicable.
5. Dilute the urine to a prescribed, constant volume (generally 1 L) with distilled water,
mix thoroughly by pouring back and forth between two containers, and collect a
prescribed portion (generally 100 mL per day from a l-L total volume) into the
container for the 10-d composite sample. Make sure that each of the 10 daily
collections is represented equally in the 10-d composite sample. (It is wise to make
duplicate, 10-d collections for each animal. Use of plastic bottles for these composite
samples will both allow freezing, if desired, and also prevent breakage.)
6. Add to the collected portion of urine an amount of concentrated HCl to ensure that the
sample is slightly acid to litmus (avoid excess). Check reaction to litmus each day,
and add acid only when necessary. RECORD the amounts of acid added. Add a few
drops of toluene to the composite sample, (as well as to the collection jugs), stopper
the composite samples, and store under refrigeration (approx. 40
C).
7. LABEL COMPLETELY:


62
a) Project number
b) Trial number
c) Animal number
d) Dates of collection
e) Experimenter
8. Inspect the apparatus and assembly for urine collection daily (or more often) to
ensure that they are clean and free-flowing.
I. Collection of feces
1. Feces must be collected daily. Follow a prescribed routine.
2. Weigh the entire excretion. Mix thoroughly, and collect a representative sample
amounting to one-tenth of the daily excretion. Record the weights of daily excretions
for each animal.
3. Into the container for the composite sample of feces add a few thymol crystals
(approx. 0.1 g per 100 g of feces collected into the vessel); mix, seal, and store under
refrigeration.
4. LABEL COMPLETELY.
IV. Analysis of samples
A. All analysis should be conducted according to recognized, accepted procedures. It
is wise for initiates to become familiar with procedures using check samples
before beginning work on experimental samples.
B. Urine. In sub-sampling urine samples for analysis, it is desirable to pour the
sample into a clean vessel - check for sediment in collection bottle and make a
quantitative transfer - and then add a (clean) magnetic stirring bar for magnetic
stirring while sub-sampling, thereby assuring suspension of any sedimentary
material. LABEL all sub-samples.
C. Feces. Composite fecal collections should be mixed thoroughly, and a
representative sub-sample collected into a weighed container suitable for drying
at 1000
C. Usually 200-g portions are selected for this determination. Dry the


63
samples to a constant weight in a forced-draft oven at 1000
C, and record loss in
weight as moisture. Such samples will lose appreciable amounts of nitrogen
and energy. These losses may be reduced by drying of the samples at 650
C, and
generally such samples are suitable for routine determinations of both nitrogen
and gross energy of feces. For precise measurements, samples of fresh feces
should be chosen for determinations of nitrogen and total energy. Alternatively,
samples of fresh feces may be dried while frozen. Dried samples should be
ground through a Wiley Mill, using at least a 2-mm screen.
D. It is a wise practice to preserve sub-samples of all feeds, feces, and urine in the
fresh state, refrigerated, until after all analyses is completed and until after the
data have been evaluated and interpreted.
Handling of Metabolic Trial Data
I. Performance
A. Body weights
B. Average daily gain
C. Average daily feed intake
D. Feed efficiency
II. Urine collection
III. Urine collection
IV. Fecal collection
V. Apparent DM Digestibility
VI. Why don’t we determine True DM digestibility?
VII. Calculations
VIII. Average values per treatment per day
A.Average values per treatment for the whole week
B. Statistical differences
1. Run SAS on your values
2. What should your model include?
3. Mean separation?


64
4. Note: When run SAS, be sure to put each animal’s values into the program.
Don’t use averages. If you need help, come see one of the instructors.
Provide means, standard errors, and p-values in a table for the following
a. Performance
1) Body weights
2) Average Daily Gains
3) Average Daily Feed Intakes
4) Feed Efficiency
b. Urine Volume
c. Fecal Production
d. Apparent Digestibility


65
Practical 16: Determination incriminating factors (Ant-nutrient factors) and Plant
secondary metabolites in feed stuffs
The nutritional value of any feed depends largely on the quality of ingredients used. This
is measured in terms of their capacity to encourage growth and maintain the animals in
good condition. Therefore, the origin of materials and/or the prior treatment to which
they are subjected influence their quality as far as the availability of nutrients that can be
used in the animal's metabolic functions. They can also affect the content of endogenous
toxic factors characteristic of each material, particulary those of vegetable origin that act
as anti-nutrients and adversely affect the organism.
There are several techniques for finding out the availability of a particular nutrient,
mainly amino-acids, and also for determining the presence of toxic substances so that
they can either be eliminated or the material rejected. This document contains the
methods for analyzing the anti-nutrients most commonly found in plant protein.
1. Rapid potenciometric method to measure urease activity in soyabean meal
This method of analysis determines residual urease in soyabean meal and its by-products
according to the Caskey-Knapp method (1944) modified by AACC (1969) and Leticia
Comar (Nutrimentos del Sureste, Merida, Yucatan, Mexico). The result is given in pH
units proportional to urease activity. Acceptable values fluctuate between 0.05 and 0.5;
lower values indicate overcooking, and higher values, undercooking.
Reagents
x Phosphate buffer solution 0.05M: dissolve 3.403 g monobasic potassium
phosphate g.r. (KH2PO4) in 100 ml recently boiled distilled water. Dissolve 4.355
g bibasic potassium phosphate g.r. (K2HPO4) in 100 ml distilled water. Combine
the two solutions, add 10 ml red phenol indicator at 0.1% and calibrate to 1,000
ml with distilled water. Adjust pH to 7.0 with an acid or strong base before using.
Kept under refrigeration, this has a useful life of 90 days.


66
x Urea buffer solution: dissolve 15 g urea g.r. in 500 ml phosphate buffer sol.
Adjust pH to 7.0 as above.
x Red phenol indicator at 0.1%: dissolve 0.1 mg phenol red in 15 ml 0.02N sodium
hydroxide sol and calibrate to 100 ml.
x Water-bath with agitator, ±0.5°C accuracy.
x pH measure with glass electrode and sufficient calomel to measure 5 ml samples
with ± 0.02 accuracy of pH units and temperature compensation.
x 20 × 150mm test-tubes with rubber stopper.
x 10 ml beakers.
Method
i. Weigh double samples of 0.2 g finely ground (without overheating) soyabean
meal and transfer to test-tubes (A and B); to tube A add 10 ml of urea buffer
solution. Stopper, shake and place in water bath for 30 min at 30 ± 0.5°C.
ii. After 5 min, add to tube B 10 ml phosphate buffer solution, stopper, shake and
place in water bath.
iii. Shake each tube every 5 minutes (6 times in 30 min).
iv. Remove tube A from water bath and decant supernatant liquid into a 10 ml beaker
and measure pH in under 3 min. Repeat process with tube B 5 min after tube A.
v. Calculate the index of urease activity as pH units resulting from the difference
between readings of tube A and tube B and determine sample quality according to
the table below:
IAU = pH tube A (sample + urea buffer) - pH tube B (sample + phosphate buffer)


67
Degree of baking pH units pH tube A pH tube B Colour
Raw 1.90 8.80 6.90 Red
Underbaked 0.80 7.60 6.80 Pink
Properly baked 0.30 7.10 6.80 Pink
Properly baked 0.08 6.98 6.90 Pale pink
Overbaked 0.03 6.93 6.90 Amber
2. Free gossypol in cottonseed meal
This anti-nutritional factor is a typical feature of cottonseed meal. Content of it must be
determined because of its detrimental effects on animals fed with it, even though fish
appear to support higher concentrations of the alkaloid due to the high protein content of
their diet. This paper describes two methods, the first for normal meal and the second for
chemically treated meals that contain di-aniline gossypol.
Reagents
x Aqueous acetone: 7 parts acetone in 3 parts distilled water (v/v).
x Aqueous acetone-aniline solution: add 0.5 ml purified aniline to 700 ml acetone
and 300 ml water. Prepare daily.
x Aqueous solution of iso-propyl alcohol: 8 parts iso-propyl alcohol + 2 parts
distilled water (v/v).
x Purified aniline: distil reagent grade aniline over a small amount of powdered
zinc, discarding the first and last 10% of distillate. Store under refrigeration in an
amber glass bottle. Stable for several months.
x Standard solution of gossypol:
a. Dissolve 25 mg pure gossypol in aniline-free acetone and transfer to a 250 ml calibrated
flask, using 100 ml acetone. Add 75 ml distilled water, calibrate with acetone and mix.


68
b. Take 50 ml of solution (a), add 100 ml pure acetone, 60 ml distilled water, mix, and
dilute to 250 ml with pure acetone. Solution (b) contains 0.02 mg of gossypol/ml and
remains stable for 24 hours in darkness.
x Mechanical agitator.
x Spectrophotometer.
x 250ml Erlenmeyer flasks.
x 25 and 250ml volumetric flasks.
x Water bath.
Method
Grind the sample so that it will pass through a 1 mm screen, taking care not to overheat.
Take approximately 1 g of the sample and add 25 ml pure acetone. Shake for a few
minutes, filter and divide the filtrate into two parts. To one part add a pellet of sodium
hydroxide and heat in a water bath for a few minutes. A yellowish extract that does not
change colour with the hydroxide indicates that the meal has not been chemically treated,
so proceed with method 1. If a deep orange-red colour appears it indicates the presence of
di-anilinegossypol and method 2 must be followed.
Method 1
Weigh 0.5–1 g of sample, depending on the amount of gossypol anticipated, into a
Erlenmeyer flask and add 2 or 3 glass beads. Add 50 ml aqueous acetone solution,
stopper the flask and agitate for on e hour. Filter, discard the first ml of filtrate and then
take two equal parts (2–10 ml, depending on the gossypol content expected) and transfer
to 25 ml volumetric flasks. Dilute one of the parts, calibrate to volume with aqueous iso-
propyl alcohol [solution (a)]. To the other part [solution (b)] add 2 ml purified aniline,
heat in water bath (90–100°C) for 30 min together with a blank containing 2 ml aniline
and a volume of aqueous acetone solution equal to the aliquot. Remove solution b and the


69
blank, add sufficient aqueous iso-propyl alcohol to make a monogeneous solution and
cool to room temperature in a bath of water. Calibrate with aqueous iso-propyl alcohol.
Read the samples in the spectrophotometer at an absorption of 400 nm. Adjust the
instrument to 0 obsorption with aqueous iso-propyl alcohol and determine the absorption
of solution (b) and the blank reagent. If the blank is below 0.022 absorption, continue
with the analysis; if not, repeat using freshly distilled aniline.
Determine the absorption of solution (b) with the blank reagent; adjust to 0 absorption.
Calculate the corrected absorption of the aliquot: absorption of solution (b) minus
absorption of solution (a). Determine the mg of free gossypol present in the solution of
sample using the calibration curve.
Method 2
Weigh 1 g of sample in a Erlenmeyer flask, add 50 ml aqueous acetone, shake and filter
as above. Add duplicate aliquots of the filtrate (2 to 5 ml, depending on the amount of
free gossypol expected) to 25 ml volumetric flasks. Calibrate one of the samples [solution
(a)] with aqueous iso-propyl alcohol and leave for 30 min before reading it on the
spectrophotometer. Calibrate the other aliquot [solution (b)] as in method (1), determine
the absorption of solutions (a) and (b) as in the method above and calculate the apparent
content of gossypol in the two solutions, using the calibration curve.
Preparation of calibration curve
Take duplicate samples of 1, 2, 3, 4, 5, 7, 8 and 10 ml of standard 0.02 mg/ml gossypol
standard and transfer to 25 ml volumetric flasks. Dilute one series [solution (a)]
calibrating with aqueous iso-propyl alcohol and determine absorption as above. To the
other series [solution (b)] add 2 ml purified aniline and proceed as above. Prepare a blank
reagent with 2 ml aniline and 10 ml aqueous acetone, heated together with the standard.
Determine absorption as in method 1 and calculate the optical density corrected for each
standard:


70
Corrected absorption = Abs. sol (b) - Abs. sol (a)
Draw the standard curve, plotting corrected absorption against gossypol concentration in
25 ml.
Calculation
Calculate % of free gossypol in normal meals:
Where: G = graph reading
W = weight of sample
V = volume of aliquot used.
For chemically treated meals:
Where: A = mg of free gossypol appearing in aliquot (a)
B = mg of free gossypol appearing in aliquot (b).
Figure 25. Determination of free gossypol in cottonseed meal
3. Determination of thioglucosides
The method described shows the approximate content of thioglucosides but does not
determine individual thioglucosides and isocyanates.
Reagents and apparatus
x 5% barium chloride solution.
x 600ml volumetric flasks.
x Vapour bath.


71
Method
i. To 10 g of meal (defatted by Soxhlet extraction) add 250 ml distilled water,
hydrolyze at 54°C for 1 h and heat at boiling point for 2 h, maintaining volume
constant.
ii. Filter, reserving the filtrate, and wash the residue three times with 50 ml hot
water, adding the liquid to the initial filtrate, and calibrate to 600 ml.
iii. Precipitate the barium sulphate by heating and adding abundant barium chloride
solution. Leave in a vapour bath for a few hours then filter.
iv. Calcine in a crucible furnace and weigh the precipitate.
Calculation
Calculate the approximate thioglucide content thus:
4. A rapid colorimetric method for the determination of mimosine (Matsumoto and
Sherman, 1951)
Mimosine is a free amino-acid very often present in certain legumes which include
Leucaena leucocephala and L. glauca, both considered as excellent sources of protein for
animal feed. Its presence limits the use of the leaves and seeds in feed for monogastric
animals since it affects thyroid function, leading to poor growth.
Reagents
x Mimosine solution. 0.1% in HCl 0.10N (recrystallized mimosine from L. glauca).
x Ferric chloride solution at 0.5% in HCl 0.1N.
x Active charcoal.
x Chlorhydric acid 0.1N.


72
x Klett-Summerson colorimeter (use spectrophotometer 535 nm) with filter N°54.
x Kitazato flask.
x Test-tubes.
x Filtration crucible.
x 250 ml beakers.
x 150 ml beakers.
x 100 ml volumetric flask.
Method
Extraction: place 1.25 g dry leucaena (kathin) leaf in a 250 ml beaker and digest with
HCl 0.1N for one hour, agitating constantly, let cool and transfer all to a 250 ml
volumetric flask. Shake well and leave until the solids settle on the bottom.
Clarification: transfer 10 ml of supernatant liquid to a 150 ml beaker with 30 mg
charcoal. Add water to bring the volume up to 25 ml, cover with a watchglass and bring
to boil for 15 min. Leave to cool and filter under vacuum in the filtration crucible; the
filtrate is collected in a test-tube placed in the Kitazato flask. Rinse the beaker and the
material in the crucible with 10 ml HCl 0.1N divided into three parts. Finally rinse the
crucible with 5 small parts of water.
Colorimetric determination: transfer the liquid obtained to a 100 ml volumetric flask, add
4 ml ferric chloride solution at 0.5% in HCl 0.1N and calibrate with water. The pH
should be between 1.5 and 2.5. Read on the colorimeter and calculate the quantity of
mimosine in a calibration curve.
Calibration curve: dissolve 0.1046 g pure mimosine in HCl 0.1N and calibrate to 100 ml
with the acid. The solution should contain 0.1% mimosine. Put 1 ml of the mimosine
solution in a 100 ml volumetric flask and add 10 ml HCl 0.1N and 4 ml ferric chloride
solution at 0.5% and calibrate with water. Read the colour intensity on the colorimeter
(spectrum 535 nm). Use distilled water as reference and repeat the process using 2, 3, 4

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