At what g force do bacteria start to pellet down a tube?

At what g force do bacteria start to pellet down a tube?

We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

I am working with C. elegans and bacteria and I want to get rid of the bacteria they eat by centrifuging the worms without centrifuging the bacteria. I am using a g-force of 600 and the bacteria sometimes still centrifuge down. What g-force would be ideal to pellet the worms but not the bacteria? Does this depend on the bacterial species?



Centrifugation is a method of separating molecules having different densities by spinning them in solution around an axis (in a centrifuge rotor) at high speed. It is one of the most useful and frequently employed techniques in the molecular biology laboratory. Centrifugation is used to collect cells, to precipitate DNA, to purify virus particles, and to distinguish subtle differences in the conformation of molecules. This chapter discusses relative centrifugal force ( g force), how to convert g force to revolutions per minute (rpm), and how to calculate the sedimentation time of a particle during centrifugation.

Prepare the Sample Tubes

You will use 1.5 mL tubes to extract the DNA samples from saliva.

To start, prepare each tube by labelling them with a permanent marker.

Even if you only have one sample, it’s good practice to label the tube clearly. For example, if the sample is from a person, you could use their initials. It’s also a good idea to mark the date of the sample.

Prepare Saline Solution

You will need salt water (saline solution) as a mouthwash to collect your cheek cells.

In a small glass or similar, mix a pinch of table salt with water. A shot glass is perfect for this. The measurements do not need to be exact. A pinch of salt for a small sip of water is a good rule of thumb.

You do not need to make much – a small sip is enough.

Why the salt water? In this protocol, the aim is to get a sample of DNA from cheek cells. Your saliva, after rinsing your mouth will naturally contain cheek cells, which will be broken open during the protocol to release the DNA. The salt, i.e. sodium chloride, is used to stabilise the DNA, once it has been released.

Rinsing your mouth

Pour the salt water into your mouth. Don’t use too much, just a small sip is enough. Rinse your inner cheeks vigorously for 30-60 seconds. When you are done, spit the saline solution into a new glass. (or, if the shot glass you used to mix the salt water is empty, you can reuse it).

The goal of this step is to loosen as many cells from your mouth as possible. You can use your teeth to gently scrape your cheeks and tongue while you are swirling the salt water around in your mouth. You can also touch your inner cheeks with your tongue. Careful to not hurt yourself – there’s no need for blood, just saliva with lots of cheek cells.

Transfer your sample into the microcentrifuge tube

For this step, you will use your saliva sample (1), the microcentrifuge tube you labelled in the beginning (2), and a transfer pipette (3).

Use the transfer pipette to transfer your saliva sample into the microcentrifuge tube. Fill it up to the 1.5 mL mark.


It is time to use the centrifuge. This will use gravitational force to concentrate the sample.

Put the centrifuge tube with your saliva sample into the centrifuge. Make sure to balance the centrifuge with another sample or with another counter weight.

If you only have one sample, the easiest way to balance the centrifuge is to fill another tube with water and use it as a balancing tube.

Using the centrifuge in an unbalanced way is dangerous and will break the device. Follow our tips for balancing a centrifuge in the manual here. In this case, for example, you could either use a second sample as a counter balance or fill up another tube with water. Tubes must always be balanced with another tube of equal weight.

Once the sample tube is balanced in the centrifuge rotor, close the lid and activate the centrifuge module. Set the speed to 4,000G and spin for 90 seconds.

Recovering the Pellet

Check the sample tube after centrifugation has finished. All the cheek cells should now be concentrated in a small white ball at the bottom of the tube (1). This is called a pellet. The remaining liquid (2), called the supernatant, should be clear.

In this step, you will remove the supernatant, so only the white pellet remains.
Check that your pellet is firmly attached to the bottom of the tube. If it is, you can carefully pour the supernatant away.

If the pellet is not firmly attached to the bottom of the tube, try spinning the sample again in the centrifuge to attach it to the bottom of the tube. If it remains loose, you can use the micropipette with a fresh tip to slowly transfer the supernatant out of the tube. You can also try using the transfer pipette you used earlier, but it might be difficult to control and could end up disturbing the pellet.

Resuspending the Pellet

You should now have a white pellet in your sample tube. It should be about the size of a matchstick head.

If your pellet is smaller than a matchstick head, you may not enough cheek cells to get a concentrated DNA sample. In that case, go back to step 2 to concentrate additional cheek cells from saliva. You can use the same sample tube and simply add more sample to the existing pellet.

Once you have a large enough pellet, you can resuspend the cells into the remaining liquid that is still in the tube. You now have a concentrated cell sample in a small volume of liquid.
Make sure the tube is closed, then mix the cells from the pellet into the liquid by flicking the tube. The cells of the pellet are now resuspended in the liquid.


In this next step, you will use the micropipette (1) to transfer the resuspended sample (2) into a 0.2mL PCR tube (3), so that you can heat it in the thermocycler.

First, set the adjustable pipette to the maximum volume of 20μl.

Make sure the pipette has a new pipette tip. Then use the pipette to transfer the cell mixture of the sample to the 0.2mL PCR tube. Carry on until the PCR tube is almost full, or until you have no sample left. Add as much of the cell mixture as possible to the PCR tube.

Labelling the PCR Tube

Finally, click the lid of the PCR tube closed and label the tube to identify the sample, similarly to the centrifuge tube.

Label the side of PCR tubes, not the lid. The PCR machine has a heated lid, so any ink on the tube lid might come off.

Heating the sample

In this step, you will use the thermocycler as a heat block to boil the cells and burst them open, to release the DNA into the solution.

Place your PCR tube with your sample cell solution in the thermocycler block.

Set up the thermocycler to heat the sample at 99°C for 10 minutes.

Mixing the Sample

After heating the sample for 10 min, we will prepare it again for centrifugation.

First, take the PCR tube out of the thermocycler block.

The block and the heated lid will still be hot, so take extra care.

Flick the PCR tube for 5 seconds to mix the sample.

Centrifuging the sample

In this step, you will spin the sample to separate the supernatant from the cell debris. Now the cells have burst thanks to the heating step, the DNA will be released from the cells and floating in the supernatant.

The molecular weight of DNA is lighter than the other cell material, like proteins and cell walls. By spinning the sample with centrifuge, we seperate the cell material from the DNA, which gives us a cleaner DNA sample.

To spin the PCR tube with your sample (3) in the Bento Lab’s microcentrifuge, you will need to use the PCR tube adapter (1) that sits in a normal microcentrifuge tube (2) and converts it to fit a PCR tube.

Remember to balance your centrifuge. So, if you are only working with one sample, prepare another PCR tube with an amount of water equivalent to your sample.

Set the centrifuge to run for 90 seconds at 8kG.

If you need help operating the Bento Lab centrifuge, check the user manual. Once you the lid is closed, select the time mode (1). Set the force (2) and time (3) before confirming.

Cleaning up the sample for storage

After centrifugation, all the cell debris has been forced to the bottom of the PCR tube (1), leaving only the DNA in the liquid supernatant (2). The supernatant should look clear, like water.

Finally, you will transfer the supernatant into a new PCR using the micropipette.

Set the micropipette to 20μL and put on a new tip. Transfer 40μL of the clear supernatant into the new PCR tube.

Be careful to avoid pipetting any cell debris into the new tube. You should only transfer the clear liquid supernatant. Avoiding any of the cell debris will reduce the chance of interference with the DNA sample.

Labeling and storage

The new tube now contains only the DNA in the liquid. It is called the template sample, and can now be further used for analysis using protocols like PCR.

Label the tube again, so that you can identify which template sample it is.

Finally, if you are not using the template sample in another protocol right away, store it in the freezer at around -20°C. This will preserve the sample.

Although DNA itself is very stable, there might still be some other proteins in the sample that will degrade it over time. The purpose of this protocol is to clean up the saliva sample as much as possible whilst retaining the DNA. Storing the sample in the freezer will slow down any reactions from left over proteins and therefore the template DNA sample will be preserved longer.

II. Increasing the effect of gravity: the centrifuge.

Many particles or cells in a liquid suspension, given time, will eventually settle at the bottom of a container due to gravity (1 x g). However, the length of time required for such separations is impractical. Other particles, extremely small in size, will not separate at all in solution, unless subjected to high centrifugal force. When a suspension is rotated at a certain speed or revolutions per minute (RPM), centrifugal force causes the particles to move radially away from the axis of rotation. The force on the particles (compared to gravity) is called Relative Centrifugal Force (RCF). For example, an RCF of 500 x g indicates that the centrifugal force applied is 500 times greater than Earth’s gravitational force. Table 1 illustrates common centrifuge classes and their applications.

Table 1. Classes of centrifuges and their applications

Table scrolls horizontally

Centrifuge Classes
Low-speed High-speed Ultra/micro-ultra
Maximum Speed (rpm x10 3 ) 10 28 100/150
Maximum RCF (x10 3 ) 7 100 800/900
Pelleting applications
Yes Yes (Yes)
Animal and plant cells Yes Yes (Yes)
Nuclei Yes Yes (Yes)
Precipitates Some Most (Yes)
Membrane fractions Some Some Yes
Ribosomes/Polysomes - - Yes
Macromolecules - - Yes
Viruses - Most Yes
Viruses - Most Yes

() = can be done but not usually used for this purpose.

Basics in Centrifugation

Centrifugation is a technique that helps to separate mixtures by applying centrifugal force. A centrifuge is a device, generally driven by an electric motor, that puts an object, e.g., a rotor, in a rotational movement around a fixed axis.

A centrifuge works by using the principle of sedimentation: Under the influence of gravitational force (g-force), substances separate according to their density. Different types of separation are known, including isopycnic, ultrafiltration, density gradient, phase separation, and pelleting.

Pelleting is the most common application for centrifuges. Here, particles are concentrated as a pellet at the bottom of the centrifuge tube and separated from the remaining solution, called supernatant. During phase separation, chemicals are converted from a matrix or an aqueous medium to a solvent (for additional chemical or molecular biological analysis). In ultrafiltration, macromolecules are purified, separated, and concentrated by using a membrane. Isopycnic centrifugation is carried out using a "self-generating" density gradient established through equilibrium sedimentation. This method concentrates the analysis matches with those of the surrounding solution. Protocols for centrifugation typically specify the relative centrifugal force (rcf) and the degree of acceleration in multiples of g (g-force). Working with the rotational speed, such as revolutions per minute (rpm), is rather imprecise.

Important definitions

In general, applications for centrifugation specify the degree of acceleration to be applied to the sample rather than specifying a specific rotational speed such as revolutions per minute. The acceleration is typically given in gravity [× g] (or multiples of x g or g-force), the standard acceleration value due to gravity at the Earth’s surface (9.81 m/s 2 ). The distinction between rpm and rcf is important, as two rotors with different diameters running at the same rotational speed (rpm) will result in different accelerations (rcf).

As the motion of the rotor is circular, the acceleration force is calculated as the product of the radius and the square of the angular velocity. Historically known as “relative centrifugal force” (rcf), this is the measurement of the acceleration applied to a sample within a circular movement. This process is measured in units of gravity
(× g).


Rotor A Rotor B
Speed 14,000 rpm 14,000 rpm
Radius 5.98 cm9.50 cm
Gravity 13,100 × g20,817 × g

As mentioned, when using
rotors with different radii for centrifugation, the same rcf
(g-force) should be used.

Both centrifuges can spin a rotor with 1.5/ 2 mL tubes at the same speed (14,000 rpm) but the acceleration applied to the samples is very different: 13,100 × g versus 20,817 × g, resulting in different results. To make life easier and to better reproduce the data, some centrifuges have buttons directly on the operating panel for automatic conversion between rpm and rcf. If your centrifuge does not have an rpm-rcf converter, you may use the formula, the rpm-rcf converter found on the homepages of centrifuge suppliers, or a nomogram for conversion. The k-factor is a parameter for the sedimentation distance in a test tube. This factor is also called clearing factor and represents the relative pelleting efficiency of a centrifugation system at maximum rotational speed. In general, the k-factor value is used to estimate the time, t (in hours), required for complete sedimentation of a sample fraction with a known sedimentation coefficient measured in s (svedberg).

A small k-factor represents a more rapid separation. The value of the k-factor is primarily determined by the rotor diameter. Compared to rpm/rcf, the usage of the k-factor has become less important for general centrifugation processes. Especially for ultracentrifugation, the k-factor is still relevant.

How to select the right centrifuge for your application

If you follow a given protocol, make sure to use the same type of rotor and apply the given relative centrifugal force (rcf) as well as the same temperature and running time. In general, the following major parameters have to be determined for a successful centrifugation run:

E: Determination of desired relative centrifugal force

F: Defined temperature during centrifugation

Fixed-angle or swing-bucket rotors

The most common rotors in laboratory centrifugation are either fixed-angle or swing-bucket rotors. Only a few applications require special rotors such as continuous-flow rotors, drum rotors, and the like. Flow-through rotors enable continuous flow collection of precipitates. These systems are used, e.g., in harvesting fermenters or for juice production in the food industry. Special customized versions, optimized for the specific application, are necessary.

Fixed-angle rotor

The obvious advantage is the lack of moving parts in the rotor. This results in lower metal stress (longer lifetime), a higher maximum g-force is possible and for many applications, faster centrifugation times can be realized. The limited capacity (less flexibility) of the fixed-angle rotor is the only drawback. The position of the pellet strongly depends on the angle of the tube, it is located from the side to the bottom of the tube when spinning. Most rotors have a 45° angle for the tubes. The larger the angle for the tubes, the tighter the pellet. Smaller rotor angles result in more spread out pellet areas.

Swing-bucket rotor

This kind of rotor is highly flexible for using different tube formats, including SBS-format plates, based on a broad range of adapter systems and a high sample capacity. The moving swing-bucket parts result in increased metal stress for the rotor and the buckets as the bucket weight places a load on the two pivots and grooves. Compared with a fixed-angle rotor, therefore, a swing-bucket rotor is limited to a lower maximum g-force, which leads to longer centrifugation times. Based on the swing-bucket principle, the pellet is located in the bottom of the tube (horizontal position of tube during the run). The recovery by the user is facilitated compared to pellets located at the side of the tube.

Methods used for Separation of Particles in Centrifugation: 3 Methods

This article throws light upon the three methods used for separation of particles in centrifugation.

The three methods are: (1) Differential Centrifugation (2) Centrifugal Elutriation and (3) Density Gradient Centrifugation. The Density Gradient Centrifugation method consists of two techniques. The two techniques are: (1) Rate Zonal Technique and (2) Isopycnic Centrifugation Technique.

Method # 1. Differential Centrifugation:

This depends upon the sedimentation rate of particles of different size and density. Centrifugations will initially sediment the largest particles.

For particles with same mass but with different densities, the one with highest density will sediment first. Particles having similar banding densities can usually be efficiently separated one from another by differential centrifugation or rate zonal method, pro­vided that there are at least 10-fold differences in their masses.

In differential centrifugation the material to be separated is divided centrifugally into number of fractions by increasing the applied centrifugal field. The centrifugal field at each step is chosen so that particular type of material sediments. Any type of particle originally present in homogenate may be found in pellet or the supernatant or both fractions, depending upon the time and speed of centrifugation and size and density of particles. At the end of each stage the pellet and supernatant are separated and pellet washed several times by re-suspension and re-centrifugation in homogenation medium.

Initially all particles of homogenate are homogenously distributed throughout the centrifuge tube. During centrifugation particles move down the centrifuge tubes at their respective sedimentation rates and start to form a pellet on the bottom of centrifuge tube. Ideally centrifugation is continued enough to pellet all the largest class of particles, the resulting supernatant then being centrifuged at a higher speed to separate medium-sized particles and so on.

However, since particles of varying sizes and densities were distributed homogenously at the commencement of centrifugation, it is evident that the pellet will not be homogenous but will contain a mixture of all the sedimented components, being enriched with fastest sedimenting particles. In the time required for complete sedimentation of heavier particles, some of the lighter and medium sized particles, originally suspended near the bottom of the tube, will also sediment and thus contami­nate the fraction.

Pure preparation of the pellet of the heaviest particle cannot be, therefore, obtained in single centrifugation step. It is only the most slowly sedimenting component of mixture remaining in the supernatant after all the larger particles have been sedimented that can be purified by single centrifugation step.

The separation achieved by differential centrifugation can be improved by repeated re-suspension and re-centrifugation under similar condition. Further centrifugation of the supernatant with gradually increasing centrifugal fields results in sedimentation of intermediate and finally the smallest and least dense particles. In spite of its inherent limitations, differential centrifugation is probably the most commonly employed method for isolation of cell organelles from, homogenized tissue.

Method # 2. Centrifugal Elutriation:

In this technique the’ separation and purification of a large variety of cells from different tissues and species can be achieved by a gentle “washing action” using an elutriator rotor. The technique is based upon the differences in the set-up in separation chamber of rotor, between the opposing centripetal liquid flow and applied centrifugal field being used to separate particles mainly on the basis of differences in their size.

The technique does not employ the density gradient and have advantage that any medium totally compatible with the particles can be used. By this separation can be achieved very quickly, giving high cell concentrations and a very good recovery yield.

Method # 3. Density Gradient Centrifugation:

There are two methods of density gradient centrifugation, the rate zonal technique and the isopycnic (iso-density or equal density) technique, and both can be used when quantitative separation of all the components of mixture of particles is required. They are also used for the determination of buoy­ant densities and for the estimation of sedimentation coefficient.

(a) Rate Zonal Technique:

Particle separation by the rate zonal technique is based upon differences in the size, shape and density of particles, the density and viscosity of the medium and the applied centrifugal field. Subcellular organelles, which have different densities but are similar in size, do not separate efficiently using this method, but sepa­ration of proteins of similar densities and differing only 3 folds in relative molecular mass can be achieved easily.

The technique involves carefully layering a sample solution on top of preformed liquid density gradient, the highest density of which does not exceed that of densest particle to be separated. The function of gradient is primarily to stabilize the liquid column in the tube against the movements resulting from conven­tional currents and secondarily to produce a gradient that helps to improve the resolu­tion of gradient.

The sample is then centrifuged until the desired degree of separation is achieved. Since the technique is time dependent, centrifugation must be terminated before any of the separated zone pellets at he bottom of tube. The technique is em­ployed for the separation of enzymes, RNA-DNA hybrids, ribosomal subunit, sub­cellular organelle, etc.

(b) Isopycnic Centrifugation Technique:

Isopycnic centrifugation depends solely upon the buyout density and not on its shape, size and time, the size of the particle affecting only the rate at which it reaches its isopycnic position in the gradient. The technique is used to separate particles of similar size but of different density. Hence soluble proteins which have very similar densities cannot be usually separated by this method, where as sub cellular organelles can be effectively separated.

The methods are a combination of sedimentation and flotation and involve layering the sam­ple on top of a density gradient that spans the whole range of the particle densities that are to be separated. The maximum density of the gradient, therefore, must always exceed the density of the densest particle. During centrifugation, sedimentation of the particle occurs until the buoyant den­sity of the particle and density of the gradient are equal.

At this point of isodensity no further sedimentation occurs, irrespective of how long centrifugation continues, because the particles are floating on the cushion of material that has density greater than their own. Isopycnic centrifugation, in contrast to the rate zonal technique, is an equilibrium method, the particle banding to form zones each at their own characteristic buoyant density.

In case when not all components in a mixture of particle are required, a gradient range can be selected in which unwanted materials will be sediment at the bottom of the tube and whole of the particles of interest will float at their respective isopynic positions. Such a technique involves a combination of both the rate zonal and isopynic approaches.

Features of Gradient Material:

There is no ideal all purpose gradient material the choice of solute depends upon the nature of the particles to be fractionated.

The gradient material should:

i. Permit the desire type of separation

iii. Be inert towards biological material and not react with the centrifuge , rotor, tubes or caps:

iv. Not absorb light at wavelengths appropriate for spectrophotometric monitoring (visible or ultraviolet range), or otherwise interfere with assaying procedures

v. Be sterilisable, non-toxic or flammable

vi. Have negligible osmotic pressure and cause minimum changes in ionic strength, pH and viscosity

vii. Be inexpensive and readily available in pure form and capable of forming a solution covering the density range needed for a particular application without overstressing the rotor

viii. Allow easy separation of the sample material from the gradient medium with loss of the sample or its activity.

Generally used gradient materials are salts of alkali metals (e.g., caesium and rubidium chloride), small neutral hydrophilic organic molecules (e.g., sucrose), hydrophilic macromolecules (e.g., pro­teins and polysaccharides), and a number of miscellaneous compounds such as colloidal silica (e.g., percoll and ludox) and non-ionic iodinated aromatic compounds (e.g., metrizamide, nycodenz and renograffin).

Sucrose solution while suffering from disadvantages of being very viscous at densities greater than 1.1 to 1.2 g cm -3 and exerting very high osmotic effects even at very low concentrations have been found to be most convenient gradient material for rate zonal separation. Glycerol is also used as gradient material especially for the separation of enzymes, or alternative media such as ficoll, metrizamide or percoll gradients may be utilized.

Non-ionic media, such as sucrose, glycerol, metrizamide, ficoll and percoll are generally considered to be gender than ionic salts like caesium chloride and potassium bromide and require a lower centrifugal fields to achieve adequate separa­tion of particles. In case of isopycnic separation, no one medium has proved satisfactory for the isolation of all type of biological particles.

Ficoll has successfully used for the separation of whole cell and subcellular organelles by rate zonal and isopycnic centrifugations. Caesium and rubidium salts are used exclusively for isopycnic separations and have been used most frequently for separa­tion of high density solutes like nucleic acids.

However, at high concentrations their high ionic strength and osmolarity tends to disrupt intra- and inter-molecular bonds. Generally, ionic media have been used for the separation of nucleic acids, proteins and viruses and non-ionic iodinated aromatic compounds, because of their increased versatility, have been used for a much wider variety of applications.

How Dying Works

After the heart stops beating, the body immediately starts turning cold. This phase is known as algor mortis, or the death chill. Each hour, the body temperature falls about 1.5 degrees Fahrenheit (0.83 degrees Celsius) until it reaches room temperature. At the same time, without circulation to keep it moving through the body, blood starts to pool and settle. Rigor mortis, or a stiffening of the body, sets in about two to six hours after death [source: Marchant, Middleton].

While the body as a whole may be dead, little things within the body are still alive. Skin cells, for example, can be viably harvested for up to 24 hours after death [source: Mims]. But some things that are still alive lead to the putrefaction, or decomposition, of the body -- we're talking about little organisms that live in the intestines.

A few days after death, these bacteria and enzymes start the process of breaking down their host. The pancreas is full of so many bacteria that it essentially digests itself [source: Macnair]. As these organisms work their way to other organs, the body becomes discolored, first turning green, then purple, then black. If you can't see the change, you'll smell it soon enough, because the bacteria create an awful-smelling gas. In addition to smelling up the room, that gas will cause the body to bloat, the eyes to bulge out of their sockets and the tongue to swell and protrude. (In rare instances, this gas has created enough pressure after a few weeks to cause decomposing pregnant women to expel the fetus in a process known as coffin birth.)

A week after death, the skin has blistered and the slightest touch could cause it to fall off. A month after death, the hair, nails and teeth will fall out. The hair and nails, by the way, while long rumored to keep growing after death, don't have any magical growth properties. They merely look bigger as the skin dries out. Internal organs and tissues have liquefied, which will swell the body until it bursts open. At that point, a skeleton remains.

Now, most of us don't see that process because the law requires that we do something with the body. There are endless possibilities: We can choose a coffin for our body or an urn for our ashes. We can be embalmed, mummified or frozen. Some cultures were rumored to engage in cannibalistic rituals of consuming the dead, while others left their dead exposed to the elements for animals to cart away. You could donate your body to science or ask for burial at sea. But unless mummified or preserved, bodies eventually disintegrate in the process described above. However, burial in a coffin slows the process tremendously even the type of soil in which you're buried can make a difference.

Disposal of a dead bod­y is largely regulated by cultural and religious beliefs. Early cultures buried the dead with their favorite possessions (and sometimes their favorite people) for the afterlife. Sometimes, warriors or servants were buried standing up, eternally ready for action. Orthodox Jews shroud their dead and bury them on the same day as death, while Buddhists believe that consciousness stays in the body for three days [source: Mims]. Hindus are cremated, because it's believed that burning releases the soul from the body, while Roman Catholics frown on cremation out of respect for the body as a symbol of human life [sources: Mims Cassell et al].

Religion and culture will always be intertwined with death, and one large area of influence relates to the ethical questions surrounding the dying process. On the next page, we'll consider some of the issues.

Practical Work for Learning

Class practical

In early studies of biology, we often focus on digestive enzymes. This can lead students to think that enzymes work only to break chemicals apart. This enzyme-catalysed synthesis offers an alternative enzyme reaction resulting in building up a new molecule.

The aim of the investigation is to prepare an extract of potato tuber, remove starch from it, and then check for its ability to catalyse the synthesis of starch from different substrates.

Lesson organisation

The enzyme extraction takes about half an hour. The extract can be stored overnight at 4 °C, but it will turn brown. To complete the investigation in one session, prepare the substrate solutions and set up the colorimeter while making the enzyme extract. Different groups could be responsible for different parts of the experiment.

Apparatus and Chemicals

For each group of students:

Access to a water bath at 25 °C

Centrifuge, 3000 rpm/ RCF 1000 g (Note 3)

2 medium-sized potatoes (Note 4)

For the class – set up by technician/ teacher:

50 cm 3 of 1% glucose-1-phosphate, for up to 8 groups (Note 1)

50 cm 3 of 1% glucose, for up to 8 groups

50 cm 3 of 1% maltose, for up to 8 groups

50 cm 3 of 1% sucrose, for up to 8 groups

Iodine solution (Note 2)

Health & Safety and Technical notes

1 Glucose-1-phosphate: Make up the solution by dissolving 0.5 g of glucose-1-phosphate in 50 cm 3 of distilled water (a 1% solution). The compound is unstable in solution, hydrolysing quite rapidly to glucose and phosphoric acid at room temperature. Prepare just before use, or store in a refrigerator (at 4 °C) for up to 24 hours. This, and the other sugar solutions, are described on the CLEAPSS Hazcard as LOW HAZARD.

2 Iodine solution: Iodine is only sparingly soluble in water (0.3 g per litre). It is usual to dissolve iodine in potassium iodide solution (KI) to make a 0.01 M solution (by tenfold dilution of a 0.1 M solution) to use as a starch test reagent. Refer to CLEAPSS Recipe card 33. For 100 cm 3 of 0.1 M solution, measure 3.0 g of potassium iodide (KI) into an appropriate beaker. Moisten the potassium iodide with a few drops of water. Measure out 2.54 g of iodine (see Hazcard 54: iodine is harmful) and add to the moistened potassium iodide. Add a small volume of water and stir. When no more iodine appears to dissolve, add some more water and stir. Repeat until all the iodine has dissolved. Pour the solution into a measuring cylinder and dilute to the final volume. If there are any bits of iodine remaining, return the solution to the beaker and leave it on a magnetic stirrer for several minutes. Add the solution to a labelled bottle and mix well.

3 Centrifuge: You need enough ‘centrifuge space’ to produce around 25 cm 3 of potato extract for each working group. A speed of just under 3000 revolutions per minute, corresponding to a relative centrifugal force (RCF, ‘g-force’) of just under 1000 g is enough to spin down starch grains.

SAFETY: The CLEAPSS Laboratory Handbook gives detailed information about the requirements of the British and International Standard (BS EN 61010-2-020) for centrifuges. Use this information to assess your school’s centrifuges if your documentation does not clearly state that they conform to this standard. The DfEE has stated that: ”Existing centrifuges not conforming to the safety requirements of this standard should not be used after 1 January 1997.” (From Safety in Science Education, HMSO, 1996, ISBN 011270915X). CLEAPSS advise that centrifuges which do not cut off the power to the rotor when the lid is raised, or have an exposed rotor, should be taken out of service. If using any centrifuge without a lock, users should not raise the lid until they can hear that the rotor has stopped.

4 Potatoes: Medium-sized potatoes are best. Large potatoes often have a poor yield of phosphorylase enzyme. Small new potatoes have such small grains of starch that they are difficult (or impossible) to spin down.


SAFETY: Ensure your centrifuge is safe to use and that students are briefed to use it correctly. Make eye protection available when iodine solution is dispensed.


a Make up a 1% solution of glucose-1-phosphate in distilled water (Note 1).

b Make up iodine solution (Note 2).

c Prepare the colorimeter by putting a known volume of water (say 3 cm 3 ) into your colorimeter cuvettes or bottle. Add 0.5 cm 3 or 1 cm 3 of iodine solution (Note 2) and swirl to mix well. Make sure this liquid is deep enough in the vessel to provide an accurate reading in your colorimeter (by comparing with a full colorimeter vessel of the same solution). Use a yellow filter, as we will be following the formation of a blue starch-iodide complex (peak of absorption 580-600 nm). Record the reading from the iodine solution as the ‘zero’ reading for this procedure. Set up ‘clean’ cuvettes to follow the reaction with 2 cm 3 of water and 0.5 or 1.0 cm 3 of iodine solution. You will add 1.0 cm 3 of starchy solution to each of these.


d Take two medium-sized potatoes, peel and cut into small pieces (Note 4). Crush with a pestle and mortar or use a blender. Add water sparingly so that the resulting mash is just liquid enough to be poured from the container.

e Pour the crushed potato quickly through a single layer of muslin or nylon stocking.

f Transfer the extract to centrifuge tubes. Spin in a bench centrifuge for 5 minutes at the highest speed to separate the starch granules (Note 3).

g After spinning, take one drop of clear liquid from the top of each tube. Test each drop on a white tile with iodine solution. If the blue colour characteristic of starch appears, centrifuge for a further 5 minutes.

h Repeat the iodine test and, if necessary, continue to centrifuge until no starch is detectable in the samples of clear liquid. Once the liquid is free of starch, carefully pour the clear liquid from each centrifuge tube into a single container. If this is to be kept overnight, stopper and refrigerate the container.

i Label 4 test tubes. Use a syringe to put 5 cm 3 of glucose-1-phosphate into a test tube. Place it in a water-bath at 25 °C. Use fresh syringes to measure 5 cm 3 of the other substrates into the other test tubes in the same water-bath. Record which substrate is in which tube.

j Use a syringe to measure four 5 cm 3 samples of the clear potato extract into each of four more test tubes, and place in the water bath at 25 °C.

k Add 5 cm 3 of potato extract from a tube in the water bath to each tube of substrate solution. Start the stopclock.

l After two minutes, take a single drop of the potato extract/ glucose-1-phosphate mixture. Mix it with a drop of iodine in one of the wells of a spotting tile. Look for a very slight colour change. If there is none, repeat at 2-minute intervals as recorded by the stopclock.

m As soon as the slightest trace of blue or grey colour appears on the tile, remove 1.0 cm 3 from the mixture into your colorimeter vessel containing 2 cm 3 of water and 0.5 cm 3 or 1.0 cm 3 of iodine solution. Mix well. Record the colorimeter reading and the time.

n Using fresh iodine solution, carry out similar measurements with the other three extract/ substrate mixtures.

o Check the amount of enzyme extract/ substrate mixture left. Arrange to take more samples at regular intervals. Try to take at least 8 readings for each extract/ substrate mix. If colorimeter readings are changing rapidly from one sample to the next with a particular substrate, concentrate on that mixture and take samples as often as possible. Then return to the other extract/ substrate mixtures.

p For each measurement, record the substrate, time, apparent colour (to your eye) and colorimeter reading.

q Convert the meter readings into starch concentrations using a calibration curve. (See associated practical on this site.)

Teaching notes

After dealing with the digestion of starch, it is appropriate to ask how starch is formed in the first place. The tissue of potato tubes is a good source of the enzyme(s) responsible for the synthesis of starch.

Digestion of starch yields first maltose and then glucose, so synthesis could be a simple reversal of this process. Alternatively, the best substrate could be one of the compounds inside cells which is closely related to simple sugars an example is glucose-1-phosphate. This demonstration shows that an anabolic pathway need not be the simple reversal of the catabolic pathway. The hydrolysis of the glucose-1-phosphate provides the energy necessary to synthesise the starch.

The starch in potato tubers is stored inside cells in large granules which sediment on centrifugation. If you examine a drop of the glucose-1-phosphate/ enzyme mixture under a microscope at the end of the investigation, you may see starch granules 4 to 10 ?m in diameter.

The ‘lag phase’ at the beginning of the process suggests that the reaction is autocatalytic. CS Hanes (1940) demonstrated this by including small amounts of soluble starch to the reaction mixture this reduced the length of the lag phase. Your students are unlikely to come up with the idea of autocatalysis on their own however it is worth discussing as an example of the way in which graphs can be used to suggest and develop hypotheses.

Health & Safety checked, May 2009


Download the student sheet Enzyme-catalysed synthesis (73 KB) with questions and answers.

Part 3: How do you choose a centrifuge?

Centrifuge speed

Centrifuges may be classified based on maximum speeds, measured as revolutions per minute (RPM). Speeds range from 0-7,500 RPM for low-speed centrifuges, all the way to 20,000 RPM or higher.

Centrifuge rotor speed is often expressed as RCF in units of gravity (x g) for various procedures. However, many centrifuges display speed as revolutions per minute (RPM), necessitating conversion to ensure the correct experimental conditions. The following formula is used to convert RPM to RCF, where R is the rotor radius (cm) and S is the speed (RPM):

Centrifuge size

Centrifuges are available as various benchtop or floor-standing models.

Floor-standing models offer greater sample capacity and can achieve high speeds. Superspeed centrifuges can achieve a maximum g-force (relative centrifugal force, RCF) of over 70,000 x g, and ultracentrifuges often used for DNA or RNA fractionation, can achieve up to 1,000,000 x g. For large-capacity, low-speed applications, low-speed centrifuges reaching approximately 7000 x g are available.

Benchtop models have a smaller footprint, and general-purpose models are ideal for a wide range of applications. There are many benchtop models available, including high-speed, microcentrifuge, clinical, and cell washer models. Clinical benchtop models and cell washers typically operate at lower speeds, and are suited to diagnostic applications, and washing debris from red blood cells.

Centrifuges for different applications

It is essential to select a centrifuge that is suited to the specific application. When purchasing a centrifuge, it is important to consider the following questions:

  • What sample volumes are you working with? For processes involving large or varying volumes, a floor-standing model with higher capacity and different rotor configurations may be the best solution.
  • Are samples temperature sensitive? If so, a centrifuge with refrigeration and temperature control options is required.
  • Will the centrifuge be used for processing clinical or blood banking samples? Cell washers or clinical models are available for these specific applications.
  • How much laboratory space is available vs the centrifuge footprint?
  • What is the maximum g-force the centrifuge is capable of generating? Low-speed centrifuges are ideal for separating whole cells, while ultracentrifuges are necessary for separating DNA and RNA.

Centrifugation speeds for cells. - (Feb/05/2013 )

What speed do you guys use to spin down your cells? Is there a rule of thumb for the different types of cells in terms of centrifugation speed?

I am having a problem with my cell count and I am not sure if the spinning down process is shearing my cells. Last night I counted my cell suspension with a hemocytometer and it came out to 1.5*10^6 cells but the FACS analysis reported a much lower number (around 10^4 cells). I know I will lose cells during the staining process and during the analysis but going from 10^6 to 10^4 is huge loss. Any advice will be appreciated.

Hm, I spin my cells down with 1300rpm for 3 min. Later during the FACS staining process I spin them down with 2400 rpm for 3 min, but be careful as this can harm the cells. Interestingly I also observe huge cell loss during the staining process although I never quantified it like you did. Do some of your cell stick to the wall of the reaction tube perhaps ?

Tabaluga on Tue Feb 5 19:12:56 2013 said:

Working with epithelia cells I use 3000 rpm for 3mins and working with HSCs the protocol calls for 1500 rpm for 5mins. I don't think my cells are sticking to the wall of the reaction tube but I don't know for sure. I don't know if it is pipetting error, cell count error or the speed of spinning.

For spinning live cells do not exceed 300 RCF (relative centrifugal forces also known as "g"), it is best to do it as low as possible, I routinely use 100 RCF for 5-10 min and it works fine for all the cancer cell lines that I work with.

When working with centrifuges, rpm is a relative measure that has no reference unless you also provide the rotor radius so Wek's 3000 rpm may in fact be lower RCF than Tabaluga's 2400, but we will never know.

Spinning too fast can cause a "smear" of cells up the wall of the tube that you may be missing when resuspending the cells.

Just looked it up, my 2400 rpm correspond to 600 g, apparently.

I will try 100, 200, and 300 RCF for 5 mins and see if there is any significant loss of cells. My 3000 rpm equals to 800 RCF.
I have actually noticed the line smear on the wall of the tube when using compensation beads but haven't really noticed it when spinning down cells.

Dear Guys,
I know this is out of ur discussion, but regarding converting RPM into G or vice verse, I just measure the radius of my centrifugal ring, not the whole machine, just the rotor radius with a ruler or something, put the data on RPM-G converter like these website:

then I got the required number in G or in RPM

great links, thanks. Some centrifuges have also got the advantage that you can switch between both values on the display, thereby detemining very quickly the relation between their rpm and g.

Just to throw in my 2 cents. In a neuro lab we spin at 90g for glia, and 160g for neurons in 15ml falcon tubes for 3 minutes.

Considering neurons are pretty small I doubt you'd need to go much faster.


  1. Baldrik

    very helpful!!! The author is just handsome !!!

  2. Darien

    Quick response, a sign of mind :)

  3. Gardalkis

    I'm sorry, but in my opinion, you are wrong. I'm sure. I propose to discuss it.

  4. Grorg

    Tell us you yourself wrote or borrowed from someone, if you yourself, then this is a rather interesting opinion

  5. Shakagul

    What audacity!

Write a message