PREPARATION OF CASEIN FROM SKIMMED MILK
INTRODUCTION
Casein is the major protein in milk. It is a phosphoprotein existing in 4 forms of kappa,
alpha, beta and gamma. This casein is a heterogenous mixture of protein compound,
containing the essential amino acids required for normal growth and development. Here,
in this experiment we attempt to isolate casein from skimmed milk to be subsequently
used in further experiments.
MATERIALS
Skimmed milk (250 ml), 1% HCl , 95% ethanol, ether, pH meter, pH paper.
PRINCIPLE: This experiment outlines the fact that a protein is least soluble at its isoelectric pH.
A protein is a chain of amino acids, some of which may contain ionizable group (acidic
or basic) at a certain pH (specific for every protein) the net positive charges balance the
negative charges i.e the overall charge on protein becomes zero. This value of pH is
called pI. The pI of casein is 4.8. To isolate casein, bring the pH down to 4.8. At this pH
the net charge on casein becomes zero. The electrostatic interaction between the water
molecules & protein would be least also the molecules of protein would be able to come
near & interact by hydrophobic interaction, aggregate to pricipitate out. HCl is used to
bring down pH to 4.8. Water is then added to remove Cl ions. Also the precipitated
protein may also contain fats which are components of milk, these are removed by
ethanol. Later ether is used to displace ethanol and water. Ether later vaporizes to give
clean and powdery casein.
PROCEDURE
1 Take 250ml skimmed milk in 1L flask/ beaker dilute it to 1L with d / w.
2 25ml 1% HCl is added with constant stirring to bring down the pH to 4.8.
3 The ppt. is allowed to settle for 15 ‘ then decanted.
4 The ppt. is washed by resuspending in 500ml d/ w. It is then allowed to settle and
decanted. Repeat this step 2/3 times.
5 The moist residue is filtered and transferred in a 250 ml beaker and a suspension of
casein is made in 100 ml ethanol. Filter.
1
6 Step 5 is repeated, 2-3 times.
7 The casein ppt. is transferred in 250 ml beaker and a suspension is made in 100 ml
ether and then filtered.
8
Step 7 is repeated 2-3 times.
9 The suspension is filtered and dried on filter paper.
10 The powder is weighed and % yield is calculated.
2
QUANTITATIVE ESTIMATION OF PROTEINS
BIURET METHOD.
Spectrophotometers and the Standard Curve
Spectrophotometers are a standard research tool used in biology and chemistry labs world
wide. Among the many uses they can follow the progress of an enzymatic reaction (as
you will do later in this course), estimate the number of bacteria in sample and detect
contaminating protein in a DNA sample. Knowing the principles behind the
spectrophotometer and how to use one is a valuable laboratory skill and you will learn the
basics of spectrophotometry in this exercise.
Learning Objectives
Upon completion of this lab we should be able to:
Define blank, light,wavelength,Absorbance,% Transmittance, serial dilution,absorption
spectrum,absorbance maximum,standard curve.How to make and use a standard curve.
Introduction to Spectrophotometers
The spectrophotometer is in principle a fairly simple instrument. But before I describe it I
need to say a couple things about light. Light is a type of electromagnetic energy and this
type of energy travels as a wave. Just like waves at the beach, electromagnetic waves
have peaks and troughs. One useful way to describe light is to measure the distance
between peaks. This distance is known as the wavelength (see Figure 1). For visible
light, these wavelengths range from about 1 380 nm (nanometers) to about 750nm. You
can see that we are talking about very small waves!
Light Traveling as a Wave
White light is made up of light of many wavelengths. Essentially a white light source
generates all visible wavelengths of light and that is a problem when using a
spectrophotometer. To get clear results, we can only measure the absorbance of one
wavelength of light at a time. Therefore, in spectrophotometer,a white light is shown on a
prism and the prism brakes up the white light into its individual wavelengths. A resulting
single wavelength of light is then selected by the operator of the spectrophotometer and
shown on a sample.
3
An experimental sample for use in a spectrophotometer usually contains a chemical in
solution or particles in a suspension which will absorb light. The amount of light the
chemical or particles absorb is measured by the spectrophotometer. That is very useful
because the amount of light absorbed can be correlated to the amount of that substance
present. Once the light is shown on a sample and some of the light is absorbed, the rest of
the light passes through the sample. The light that passes through is collected and
measured by a phototube and the results are displayed on a scale. The scale on a
spectrophotometer is something else that deserves mention. Look at a spectrophotometer.
You will notice that there are actually two scales: Absorbance and % Transmittance.
No, these weren’t placed here specifically to confuse you. It turns out that some
experiments or procedures work better with one scale or the other. But lets bypass that
and move on to what the scales actually measure. % Transmittance is a scale that
measures the amount of light that shines through a solution. For example, if 75% of the
light shining through a sample makes it all the way through, that would be 75%
Transmittance. If a sample completely blocked the light shining on it, what would be the
% Transmittance? If you answered 0% Transmittance, you have the idea. How about
Absorbance? Absorbance measures the amount of light absorbed by a sample (i.e.
blocked from passing through). That sounds like the opposite of the definition of %
Transmittance and it is! To further confirm this, look at the scales. They run in the
opposite directions. The Absorbance scale as you can see is not linear like the
%Transmittance scale but exponential, starting at zero and ending at infinity. This allows
for great sensitivity at the low end of the scale but poor sensitivity at the high end. For
this reason Absorbance readings above about 1.0 are usually not used. Also note that the
Absorbance scale is only a relative scale and therefore has no units. Before you go on
with this exercise, there are some things about the absorbing molecule (or particle) and
the amount of light absorbed by that molecule (or particle) that you should know. The
amount of light absorbed depends on several factors.
1. The higher the concentration of the absorbing molecule, the more
light is absorbed.
2. The longer the distance that the light travels through the sample,
the greater the absorbance. This factor is held constant because all of the
samples are measured in the same sized test tube.
3. A substance that absorbs a light of one specific wavelength very
strongly may absorb light of another wavelength only weakly. For this
reason, different wavelengths of light are used to measure the concentration
of various substances.
One more thing before you tackle the experiments. When an object or a solution is
illuminated, some wavelengths of light are absorbed and some are reflected. The color of
an object or solution that you see is composed of the wavelengths of light that are being
reflected. The colors that you don’t see are the wavelengths of light that are being
absorbed.
The absorption spectrum of Biuret reagent
4
This blue colored reagent is commonly used to stain proteins. Proteins by themselves are
usually colorless and therefore it is difficult to measure the presence of protein without a
special kind of spectrophotometer (which happens to be very expensive). An easy (and
inexpensive!) way to remedy this problem is to stain the protein with Biuret reagent and
measure the amount of absorption of the Biuret reagent.
However you should know what wavelength of light is best absorbed by theBiuret
reagent.
Blanks -You will be using Blanks in these experiments. Blanks are useful when there are
other substances in the experimental tube besides the substance you are trying to
measure. Since those other substances are not the chemical that you are trying to
measure, they often interfere with the absorbance reading of the chemical of interest.
Remember: you are interested only in the Absorbance due to the substance you want to
measure.
How do you deal with this problem? One way would be to remove the offending
substances, but this is usually not practical and often the substances are necessary in the
experiment. A much more suitable way to deal with this problem is to exclude these other
substances from our spectrophotometer reading without removing them from the
experimental tube. The way to do that is to use a blank. A blank contains all the
substances (or substance) in the experimental tube except the substance that is being
measured. Then, before reading your experimental tube, you place the blank tube in the
spectrophotometer, set the spec. to “0” Absorbance (or 100% Transmittance), then read
the experimental tube. What you have done, is to set the spec. to read only the absorbance
that is due to the substance that you are interested in!
How does biuret reagent cause a color change with proteins?
The reagent used in the Biuret Test is a solution of copper sulfate (CuSO4) and potassium
hydroxide (KOH). The KOH is there to raise the pH of the solution to alkaline levels; the
crucial component is the copper (II) ion from the CuSO4.
When peptide bonds are present in this alkaline solution, the copper (II) ions will form a
coordination complex with four nitrogen atoms involved in peptide bonds, as described in
the figure below.
5
In this figure, the nitrogens on the left are adjacent in the sequence of one peptide, and
the nitrogens on the right are adjacent in the sequence of another peptide (or another
section of the same peptide). As you can see, the longer a peptide, the more of these
complexes you can form.
Copper Sulfate solution is a blue color, but when the copper (II) ions are coordinated with
the nitrogen atoms of these peptide bonds, the color of the solution changes from blue to
violet. This color change is dependent on the number of peptide bonds in the solution, so
the more protein, the more intense the change. When the peptides are very short, the
solution turns a pink color, rather than violet.
Preparation of reagents
A .Biuret reagentCuSO45H2O
0.3 gm
NaK Tartrate
0.9 gm
NaOH
0.8 gm
KI
0.5 gm
Dissolve separately in d. w. mix and makeup the volume to 100 ml.
B. Standard Protein- 5 mg in 0 .2 N NaOH
METHOD
3ml Biuret reagent is added to 2ml of protein solution. Mix. And Warm at 37 0C
for 10 min. Cool and read absorption at 540nm.
OBSERVATION
Tube
no.
Casein
ml
Water
ml
Biuret
reagent
O.D. at 540
nm
Amount of
protein
1
2
3
4
5
6
7
6
unknown
1
2
3
7
Making and Using a Standard Curve
Serial dilution A serial dilution is a set of dilutions in which the important reagent is
present in a regularly decreasing / increasing concentration.
What is a standard curve?
As the concentration of a light absorbing chemical in a solution increases, the absorbance
of the solution increases. If we make a serial dilution of the chemical in a solution and
then take absorbance readings, we can construct a graph which shows the relationship
between concentration and absorbance. That is a standard curve and the standard curve
has many applications in the biology and chemistry lab when the concentration of a
solute in a sample is determined.
Making of a standard curve
Make a “best fit” line graph of your results by hand on graph paper. The Y” axis
should be the Absorbance scale and the “X” axis will be the Concentration scale. This
graph of a standard curve will be turned in with the exercise. Determination of the
concentration of an unknown sample is simple once you have a graph of a standard curve
for that solute.
1) Locate the absorption value of the unknown on the Absorbance
scale( “X” axis)
2) Draw a horizontal line at this value across the graph until it
intersects with the standard curve
3) At this point, draw a line vertically down until it intersects the
Concentration scale (“Y” axis)
4) The concentration value at this point will be the solute
concentration of the unknown solution.
8
Complication
All assays have limits. Amounts of substance below some minimum will be undetectable.
Beyond some maximum amount or concentration an assay becomes saturated, that is,
increases in amount or concentration do not affect absorbance. We generally try to work
within the linear range of an assay, that is, where absorbance is directly proportional to
concentration. Ideally, we would set up standards that encompass the entire useful range
of an assay. That is, we optimize the range of the assay.
Often a sample is so concentrated that when you assay the prescribed volume of sample
the result is off scale – the assay reagent is saturated. The solution then is to dilute the
sample. For example, if the volume of each standard or sample is 1 ml, and 1 ml of your
unknown gives a result that is off scale, you can add 0.1 ml sample to a test tube along
with 0.9 ml buffer. If you read a concentration from the standard curve, then multiply the
result by 10 to get the actual concentration in the sample. If you read an amount from the
standard curve then simply divide that amount by 0.1 ml to get your concentration.
When samples are so concentrated that you cannot pippet a small enough amount
accurately, you may have to conduct serial dilutions.
9
The Lowry Assay
References:
O.H. Lowry, N.J. Rosebrough, A.L. Farr and R.J. Randall (1951) J. Biol. Chem. 193:
265. (The original method)
How does it work?
●
●
The first step is a Biuret reaction which reduces Cu+2 to Cu+1
The second reaction uses Cu+1 to reduce the Folin-Ciocalteu reagent
(phosphomolybdate and phosphotungstate). This is detectable in the range of 500
to 750 nm
Detection Limitations
●
2-100 µg
Advantages
●
●
●
●
Sensitive over a wide range
The most commonly referenced procedure for protein determination
Can be performed at room temperature
10-20 times more sensitive than UV detection
10
●
Can be performed in a microplate format
Disadvantages
Many substances interfere with the assay
Alkaline copper reagent is laborious to prepare and will develop carbonate scales
over storage which interfere with optical activity, thus it must be prepared fresh
daily
● Takes a considerable amount of time to perform
● The assay is photosensitive, so illumination during the assay must be kept
consistent for all samples
● Amount of color varies with different proteins
●
●
General Considerations
Some researchers have reported that repeated assays in the same cuvettes cause
them to be etched
● Many chemical distributors sell a modified Lowry assay that is more stable and
sensitive than homemade versions
● Since reduced copper is detected in the procedure, make sure that the distilled
water used in the procedure is fed from plastic lines and not copper lines. In
general water from 18 mega ohm water polishers is satisfactory
● Variation in the content of tyrosine and tryptophan residues will influence the
assay.
●
MARERIALS
(1)
(2)
(3)
(4)
(5)
(6)
Alkaline Na2CO3, 2% solution.
CuSO4, 2% + Na-K tartarate, 1% solution.
Alkalie solution – prepared by mixing 100ml of (1) and 2ml of (2)
Folin reagent. It is diluted with equal amount of water just before use.
Standard BSA. 0.2mg / ml
Unknown sample.
Procedure:
●
●
●
●
●
●
●
Add samples containing up to 100 µg of protein
Bring all tubes to 1 ml total volume with water.
Prepare the Assay Mix and diluted Folin-Ciocalteu reagent.
To each tube add 5 ml of assay mix and thoroughly vortex.
Incubate tubes at room temperature for 10 min.
Add 0.5 ml of diluted Folin-Ciocalteu reagent. Vortex immediately.
Incubate at room temperature for 30 min.
11
●
Vortex the tubes, zero the spectrophotometer with the blank and measure
absorbance at 660 nm (or other appropriate wavelength). The data from the
standard curve are usually linear enough that a straight-line interpolation can be
used to determine the concentration of unknowns.
OBSERVATION
Tube.no. B.S.A.
water
Alkaline
Reagent
Foline
Reagent
O.D.at
600nm
1
2
3
4
5
6
7
8
unknown
1
2
3
12
Discussion
The Lowry method relies on two different reactions. The first is the formation of a
copper ion complex with amide bonds, forming reduced copper in alkaline solutions.
This is called a "Biuret" chromophore. The second is the reduction of Folin-Ciocalteu
reagent (phosphomolybdate and phosphotungstate) by tyrosine and tryptophan residues.
The reduced Folin-Ciocalteu reagent is blue and thus detectable with a spectrophotometer
in the range of 500-750 nm. The Biuret reaction itself is not all that sensitive. Using the
Folin-Ciocalteu reagent to detect reduced copper makes the assay nearly 100 times more
sensitive than the Biuret reaction alone.
The assay is relatively sensitive, but takes more time than other assays and is susceptible
to many interfering compounds. The following substances are known to interfere with
the Lowry assay: detergents, carbohydrates, glycerol, Tricine, EDTA, Tris, potassium
compounds, sulfhydryl compounds, disulfide compounds, magnesium and calcium. Most
of these interfering substances are commonly used in buffers for preparing proteins. This
is one of the major limitations of the assay. The Lowry assay is sensitive to variations in
the content of tyrosine and tryptophan residues. If the protein you are assaying has an
unusual content of these residues, an appropriate substitute standard is required. The
standard curve is linear in the 1 to 100 ug protein region. The absorbance can be read in
the region of 500 to 750 nm. Most researchers use 660 nm, but other wavelengths also
work and may reduce the effects of contamination (e.g. chlorophyll in plant samples
interferes at 660 nm, but not at 750 nm).
Typical Standard Curve for a Lowry Assay
13
CALCULATION AND RESULT
Bradford protein assay
The Bradford assay is very fast and uses about the same amount of protein as the Lowry
assay. It is fairly accurate and samples that are out of range can be retested within
minutes. The Bradford is recommended for general use, especially for determining
protein content of cell fractions and assesing protein concentrations for gel
electrophoresis.
Principle
The assay is based on the observation that the absorbance maximum for an acidic
solution of Coomassie Brilliant Blue G-250 shifts from 465 nm to 595 nm when binding
to protein occurs. Both hydrophobic and ionic interactions stabilize the anionic form of
the dye, causing a visible color change. The assay is useful since the extinction
coefficient of a dye-albumin complex solution is constant over a 10-fold concentration
range.
Equipment
A visible light spectrophotometer is needed, with maximum transmission in the region of
595 nm, on the border of the visible spectrum (no special lamp or filter usually needed).
Glass or polystyrene (cheap) cuvettes may be used, however the color reagent stains both.
Disposable cuvettes are recommended.
14
Reagents
1. Bradford reagent: Dissolve 100 mg Coomassie Brilliant Blue G-250 in 50 ml 95%
ethanol, add 100 ml 85% (w/v) phosphoric acid. Dilute to 1 liter when the dye has
completely dissolved, and filter through Whatman #1 paper just before use.
2. (Optional) 1 M NaOH (to be used if samples are not readily soluble in the color
reagent).
The Bradford reagent should be a light brown in color. Filtration may have to be repeated
to rid the reagent of blue components. The Bio-Rad concentrate is expensive, but the lots
of dye used have apparently been screened for maximum effectiveness. "Homemade"
reagent works quite well but is usually not as sensitive as the Bio-Rad product.
Assay
1. Warm up the spectrophotometer before use.
2. Dilute unknowns if necessary to obtain between 5 and 100 µg protein in at least
one assay tube containing 100 µl sample
3. If desired, add an equal volume of 1 M NaOH to each sample and vortex (see
Comments below). Add NaOH to standards as well if this option is used.
4. Prepare standards containing a range of 5 to 100 micrograms protein (albumin or
gamma globulin are recommended) in 100 µl volume. Add 5 ml dye reagent and
incubate 5 min.
5. Measure the absorbance at 595 nm.
Analysis
Prepare a standard curve of absorbance versus micrograms protein and determine
amounts from the curve. Determine concentrations of original samples from the amount
protein, volume/sample, and dilution factor, if any.
Preparation of standard curve
Sr.no
BSA
Standard
O µl
20
40
60
80
100
1
2
3
4
5
6
Unknown
1
50
water
Coomassie Dye
100 µl
80
60
40
20
0
5
5
5
5
5
5
50
5 ml
O.D.at 540nm
ml
ml
ml
ml
ml
ml
15
2
100
0
5 ml
Comments
The dye reagent reacts primarily with arginine residues and less so with histidine, lysine,
tyrosine, tryptophan, and phenylalanine residues. Obviously, the assay is less accurate for
basic or acidic proteins. The Bradford assay is rather sensitive to bovine serum albumin,
more so than "average" proteins, by about a factor of two. Immunoglogin G (IgG gamma globulin) is the preferred protein standard. The addition of 1 M NaOH was
suggested by Stoscheck (1990) to allow the solubilization of membrane proteins and
reduce the protein-to-protein variation in color yield.
References
●
Bradford, MM. A rapid and sensitive for the quantitation of microgram quantitites
of protein utilizing the principle of protein-dye binding. Analytical Biochemistry
72: 248-254. 1976.
Ultraviolet Absorbance of Aromatic Amino Acids
Reference: E. Layne. Methods in Enzymology 3: 447 (1957)
How does it work?
●
Monitors the absorbance of aromatic amino acids, tyrosine and tryptophan or if
the wavelength is lowered, the absorbance of the peptide bond. Higher order
structure in the proteins will influence the absorption
Detection Limitations
●
20 µg to 3 mg
Advantages
●
●
●
●
Quick
Sample can be recovered
Useful for estimation of protein before using a more accurate method
Well suited for identifying protein in column fractions
16
Disadvantages
Highly susceptible to contamination by buffers, biological materials and salts
Protein amino acid composition is extremely important, thus the choice of a
standard is very difficult, especially for purified proteins
● Absorbance is heavily influence by pH and ionic strength of the solution.
●
●
General Considerations
●
This is often used to estimate protein concentration prior to a more sensitive
method so the protein can be diluted to the correct range
Quantitative Procedure
Zero the spectrophotometer with a buffer blank
Make a standard curve using your standard of choice in the expected
concentration range, using the same buffer that your unknown sample is in.
● Take the absorbance values at 280 nm in a quartz cuvette
● Place sample into quartz cuvette (make sure concentration is in the range of 20 µg
to 3 mg
● Take absorbance at 280 nm
●
●
Estimation Procedure
●
●
●
●
●
Zero spectrophotometer to water (or buffer)
Take the absorbance at 280 nm in a quartz cuvette
Change wavelength to 260 nm and zero with water (or buffer)
Take absorption at 260 nm in a quartz cuvette
Use the following equation to estimate the protein concentration
[Protein] (mg/mL) = 1.55*A280 - 0.76*A260
Preparation of standard curve
Sr. No.
1
Standard protein
0 ml
water
1.0 ml
2
0.2
0.8
3
0.4
0.6
4
0.6
04
5
0.8
0.2
O.D.at280nm
O.D.at 260nm
17
6
Unkown
1
1,0
0
Auto zero with
blank
2
1.0 ml
Discussion
Determination of protein concentration by ultraviolet absorption (260 to 280 nm) depends
on the presence of aromatic amino acids in proteins. Tyrosine and tryptophan absorb at
approximately 280 nm. Higher orders of protein structure also may absorb UV light or
modify the molar absorptivities of tyrosine and tryptophan and thus the UV detection is
highly sensitive to pH and ionic strength at which measurement is taken. Many other
cellular components, and particularly nucleic acids, also absorb UV light. The ratio of
A280/A260 is often used as a criterion of the purity of protein or nucleic acid samples
during their purification. The real advantages of this method of determining protein
concentration are that the sample is not destroyed and that it is very rapid. Although
different proteins will have different amino acid compositions and thus different molar
absorptivities, this method can be very accurate when comparing different solutions of
the same protein.
To make an accurate determination of protein concentration, you will have to produce a
standard curve (A280) with known amounts of purified protein. You will also have to
provide a blank that is appropriate for the sample and contains the same concentrations of
buffer and salts as the sample. It is often convenient to dialyze the sample and measure
the absorbance of the retentate (still in the dialysis sack) using the dialysate as the blank.
Care must be taken to use quartz cuvettes, since glass absorbs UV light. A handy
equation to estimate protein concentration that is often used is
[Protein] (mg/mL) = 1.55*A280 - 0.76*A260
However, it is also a good idea to always use a standard curve and suggested that you
evaluate the agreement of the results using the above equation with results using a
standard curve.
This method is the least sensitive of the methods discussed here. For increased
sensitivity, the wavelength can be lowered to the range of 210 to 225 nm. This measures
the amide bond in proteins. However it is much more subject to interference from many
more biological components and compounds used to make buffer solutions.
18
If you don't know what the protein concentration of an unknown sample is likely to be,
the ultraviolet method might be a good starting point. Prepare a standard curve for the
absorbance at 280 and 260 nm. After you have the data for the standard curve, rezero the
spectrophotometer with water. Place your samples into a dry 1 mL quartz cuvette and
read the absorbance. If the A280 of your unknown sample is less than 2, you should
probably not dilute your sample further. If the absorbance is >2, dilution will be required.
When you are finished with the first measurement, the unknown can be returned to its
original tube with minimal loss.
Estimation of Inorganic Phosphate
Fiske and Subbarow method
Protein’s phosphorus is estimated after converting it to inorganic phosphate(Pi),which is
done by hydrolyzing the phosphorus, by an anhydrous acid. Following the release of the
phosphorus it is estimated by the Fiske and Subbarow method.
Of the many methods available for the estimation of phosphate, that of Fiske and Subba
Row is one of the simplest and most widely used. In certain cases, however, it is
convenient to use procedures in which the phosphomolybdate complex is extracted with
an organic solvent such as isobutanol, thus avoiding the interference due to colored
substances, citrates, oxalates, buffers etc.
In the Fiske and Subba Row procedure the oxtra-labile compounds are estimated as
inorganic phosphate because of the relatively high acid concentration (pH 0.65) and
because molybdate accelerates the hydrolysis of some organic phosphates. By measuring
19
the color immediately after adding the reagents, however, Fiske and Subba Row were
able to estimate phosphsocreatine.
Hydrolysis of Casein for the estimation of phosphate.
Weigh 100mg casein(prepared from milk) .
Transfer it into a glass ampoule.
Fill the ampoule with 6N HCl.
Get the ampoules sealed by the Glass Blower.
Heat on a boiling water bath for 18 hours.
Reagents:5N Sulfuric acid.
2.5 % Ammonium molybdate.
Reducing agent. This may be prepared in the powdered form and dissolved before use.
The solution deteriorates slowly and should not be used after more than a week. The
powdered reagent is prepared by mixing thoroughly 0.2g. of 1-amino-n-naphthol sulfonic
acid with 1.2 g. of sodium bisulfate and 1.2gm.of sodium sulfite. For use of 0.25g. is
measured with a small spoon and dissolved in 10 ml of water.
Standard Solution.1.3613g of analytically pure KH2PO is dissolved in 1000 ml of water, a
few drops of chloroform are added, and the solution is stored in the refrigerator. For use
it is diluted 1: 10, so that 1 ml corresponds to 1 micromole of phosphorus.
Unknown phosphate sample
Break open the glass ampoule containing hydrolysed casein sample. Measure and note
down it’s volume. Use 0.1, 0.2, 0.5, ml for the estimation of phosphate.
PROCEDURE:The standard and unknowns should contain from 0.1 to 1 micromole of phosphate. One
milliliter of sulfuric acid is added followed by 1ml. Of moldboard. After mixing 0.1 ml.
of reducing solution is added. The volume is made up to 10ml. After mixing again,
absorbency at 660mu. Is measured after 10 minutes.
PREPARATION OF SOLUTIONS
1. 5NH2SO4
2. Amm. Molybdate
3. Reducing Reagent
13.9 ml / 100 ml of conc. H2SO4
1gm / 40ml
1 amino-2-naphthol 4-sulfonic acid.
0.2 gm
Sod. Bisulfate
1.2 gm
20
Sod. Sulfite
1.2 gm
Dissolve in about 100 ml d.w. before use. (prepare fresh )
OBSERVATION
Tube Standard
water 5NH2SO4 Molybdate Reducing Warter O.D.at
no.
phosphate
reagent
reagent
600nm
1
2
3
4
4
6
7
UNKNOWN
1
2
3
21