Key takeaways

• RCF = 1.118 × 10⁻⁵ × r × N², where r is the rotor radius in centimetres and N is speed in rpm.
• To go the other way: N = √( RCF ÷ (1.118 × 10⁻⁵ × r) ). This is the direction most people actually need, since protocols specify ×g.
• The constant is fixed physics. It does not vary between centrifuge models — what varies is the rotor radius, and that is why the same rpm gives different forces on different machines.
• Radius matters more than most people expect: RCF scales linearly with r, so a rotor twice the radius doubles the force at identical rpm.
• Speed matters even more: RCF scales with the square of rpm, so doubling the speed quadruples the force.
• Use r_max for pelleting unless a protocol states otherwise, and record which radius you used — r_min and r_max can differ by a factor of two in the same rotor.
• Always publish and follow protocols in ×g, not rpm, or the method cannot be reproduced on different equipment.

Contents

• Why rpm alone is not enough
• The conversion formula
• Converting g-force back to rpm
• Worked examples
• Which radius: r_min, r_avg or r_max
• Quick reference table
• Rotor type and practical factors
• Common errors
• Sample prep reagents
• Frequently asked questions

Reagents for spin-based sample prep

Centrifugation is a step rather than an endpoint. The reagents below cover the workflows where the spin settings above matter most — lysate clarification, exosome isolation and post-spin quantification.

Lysis

GenieLyse RIPA Lysis Buffer

Lysis bufferCells & tissue

Lysates need a clarifying spin to pellet debris before assay — typically a high-speed, short, cold step.

View product →

Exosomes

Exosome Isolation Kit (Precipitation)

PrecipitationSerum, plasma, media

A polymer-based alternative to differential ultracentrifugation, using standard benchtop speeds.

View product →

Affinity

Exosome Isolation Affinity Purification

AffinitySerum, plasma, media

Affinity capture where ultracentrifuge access is limited or higher purity is needed.

View product →

Bradford

Bradford Protein Assay

ColorimetricProtein lysates

Quantifies protein in the clarified supernatant; incompatible with detergents above their limits.

View product →

Total protein

Total Protein Assay Kit

ColorimetricProtein lysates

An alternative total protein readout for normalising downstream assays after clarification.

View product →

Viability

Propidium Iodide Staining Solution

DNA stainFlow cytometry

Checks viability after processing — excessive g-force damages cells and shows up here.

View product →

Why rpm alone is not enough

Revolutions per minute describes how fast the rotor turns. It says nothing about the force acting on the sample, because that depends on how far the sample sits from the axis of rotation.

Two centrifuges running at an identical 5,000 rpm will subject samples to quite different forces if their rotors differ in size. A protocol written as “5,000 rpm for 10 minutes” is therefore not reproducible on other equipment, whereas “3,000 × g for 10 minutes” is. Relative centrifugal force is expressed as a multiple of Earth’s gravitational acceleration, which is why it is written ×g and carries no other units.

The conversion formula

To convert speed into force:

RCF = 1.118 × 10⁻⁵ × r × N²

• RCF is relative centrifugal force, in ×g.
• r is the rotational radius in centimetres — the commonest single mistake is entering millimetres or inches.
• N is rotor speed in revolutions per minute.

If the radius is measured in millimetres instead, the constant becomes 1.118 × 10⁻⁶. The two forms are equivalent; mixing them produces an answer wrong by a factor of ten.

The constant itself is derived from physics — it packages the conversion between angular velocity in rpm and acceleration relative to g. It is the same for every centrifuge ever made. If a manufacturer’s documentation appears to give a different conversion, it is quoting a value for a specific rotor radius, not a different constant.

Converting g-force back to rpm

This is the direction most often needed, because protocols specify ×g while many centrifuge dials are set in rpm. Rearranging gives:

N = √( RCF ÷ (1.118 × 10⁻⁵ × r) )

In practice: divide the required RCF by the constant multiplied by your rotor radius in centimetres, then take the square root. Because of that square root, a modest change in required force translates into a smaller change in rpm — quadrupling the force needs only a doubling of speed.

Worked examples

Speed to force

A rotor of radius 10 cm running at 10,000 rpm:

• RCF = 1.118 × 10⁻⁵ × 10 × 10,000²
• = 1.118 × 10⁻⁵ × 10 × 1 × 10⁸
• = 11,180 × g

Force to speed

A protocol requires 3,000 × g and your rotor radius is 8 cm:

• N = √( 3,000 ÷ (1.118 × 10⁻⁵ × 8) )
• = √( 3,000 ÷ 8.944 × 10⁻⁵ )
• = √( 33,542,000 ) ≈ 5,792 rpm

Round to a setting your centrifuge can actually hold, and record both figures in your notes.

Which radius: r_min, r_avg or r_max

This is the detail that causes most real-world discrepancies, and it is absent from most conversion guides. A tube is not a point — it occupies a range of radii, so the sample experiences a range of forces.

The gap is not trivial. In a microcentrifuge rotor, r_min might be 4 cm and r_max 8 cm — a two-fold difference in force between the top and bottom of the same tube at the same speed. Two labs following one protocol can therefore differ two-fold simply by choosing different radii.

Manufacturers publish these figures for each rotor. Use r_max unless the protocol says otherwise, and state which you used when writing methods.

Quick reference table

Relative centrifugal force in ×g, calculated with the formula above:

Read across to see how strongly radius matters at fixed speed, and down to see the squared effect of speed. Use the table for orientation and the formula for your actual rotor.

Rotor type and practical factors

• Fixed-angle rotors hold tubes at a set angle. The effective radius changes little during the run, and pellets form against the tube wall as well as the bottom.
• Swinging-bucket rotors let tubes pivot to horizontal at speed, so r_max during the run differs from the value at rest. Pellets form flat at the tube bottom, which is preferable for gradients and for resuspending cleanly.
• Acceleration and braking are not captured by either number. Density gradients require slow acceleration and braking off, or the separation is destroyed regardless of the correct RCF.
• Temperature matters for labile samples and for density-based separations, where medium density is temperature dependent.
• Balance is a safety requirement as well as a technical one: opposing tubes should be matched by mass, not by eye.

Common errors

Frequently asked questions