Does Centrifuging Cause Hemolysis? What the Evidence Shows

Hemolysis is one of the top reasons clinical specimens get rejected. Most labs assume the centrifuge is the reason why, but a single, properly run spin adds only a small amount of free hemoglobin to a specimen—far below the level at which a sample gets flagged as hemolyzed. Most hemolysis is set in motion before the tube ever reaches the rotor, during the draw and during transport, where the collection method itself is a leading driver of hemolyzed samples (CDC Laboratory Medicine Best Practices review). The spin step can contribute, but only through a short and preventable list of factors, and re-spinning the same tube is the one that matters most.

This piece separates what the spin step actually does from what gets blamed on it and shows how to remove the few variables a lab controls during centrifugation.

What hemolysis is, and why it skews results

Hemolysis is the rupture of red blood cells, which releases free hemoglobin and other intracellular components into the serum or plasma. The problem is not cosmetic. Several analytes sit at much higher concentrations inside red cells than in plasma, so when cells break open those analytes read falsely high. Potassium is the clearest example, along with lactate dehydrogenase (LDH) and aspartate aminotransferase (AST). A hemolyzed potassium result can read high enough to suggest a clinical emergency the patient does not have, which is why labs reject these samples rather than report them.

Hemolysis is also very common. In a review of specimen quality, hemolysis was the single most frequent reason samples were judged unsuitable, accounting for roughly 40 to 70 percent of all rejected specimens (Lippi et al., 2008). A 2023 systematic review and meta-analysis framed it slightly differently, ranking clotting first at about 32 percent of rejections and hemolysis second at about 23 percent (Getawa et al., 2023). Either way, it is one of the top two preanalytical reasons a sample never gets reported, so the question of what causes it is worth getting right.

Does centrifuging blood cause hemolysis?

For a single, properly performed spin, centrifuging blood causes very little hemolysis. In experiments on human whole blood, a single spin left free hemoglobin no higher than in blood left to settle without spinning. This result was seen in samples centrifuged up to 16,000 ×g, far above any clinical protocol. Measured free hemoglobin increased beyond the control only after the sample was subjected to consecutive re-spinning (Mancuso et al., 2018). A separate study measured a rise in free hemoglobin from one centrifugation step, at a force of 900 ×g and found only a small rise over its unspun baseline: from about 61 to 79 mg/L (p = 0.002) (Wiegmann et al., 2017).

Now to put those numbers in context with the thresholds used in a lab. Most labs flag a specimen as hemolyzed when free hemoglobin reaches roughly 0.3 g/L (300 mg/L), where CLSI places the boundary of a high degree of hemolysis (CLSI GP44-A4 / PRE04), though exact flagging thresholds vary by analyte and analyzer. A single, properly run spin lands well under that. So the honest answer is that the spin contributes, but a correctly performed single centrifugation does not, on its own, push a normal sample across the line into rejection. What changes that is re-spinning the sample.

One caveat belongs here, because it is the kind of detail that gets lost. Those measurements come from research experiments on whole blood, not from routine clinical collection tubes. They tell us the direction and the rough size of the effect, not the clinical cutoff. For that, the standard is the better anchor.

CLSI guidance on specimen handling lists the recognized in-vitro causes of hemolysis, and routine single centrifugation is not among them. The listed causes sit on the collection and handling side: difficult venipuncture, small-gauge needles, especially winged sets on evacuated tubes, drawing through a catheter, and rough handling such as rimming a clot (CLSI GP44-A4 / PRE04). When the standard names the causes and the spin step is absent from the list, that tells you where to look first.

Which centrifuge-related factors genuinely contribute?

A short list, in rough order of how often it actually matters:

• Re-spinning the same specimen. Every additional spin of the same tube adds to the total free hemoglobin, and this is the parameter the data ties most directly to hemolysis. When a sample needs more plasma or a cleaner separation, spinning it a second time is the habit most worth breaking. Usually needing more plasma is patient specific and in most cases would require an additional blood draw. If you are consistently producing poorly separated blood tubes, it may be time to ensure your centrifuge is working properly.
• Exceeding the tube’s maximum rated force. Blood collection tubes carry a maximum relative centrifugal force in their instructions for use. Spinning above it risks tube breakage, which means a lost sample plus an aerosol and biosafety exposure for the tech at the rotor. Follow the tube manufacturer’s stated maximum (for example, the BD Vacutainer or Greiner VACUETTE instructions for use) and do not exceed it to save time.
• Rotor type for gel separator tubes. Gel barrier tubes are validated for swing-out rotors, where the tube hangs vertical during the spin and the gel forms a clean horizontal barrier. Fixed-angle rotors hold the tube at an angle and are not recommended for gel tubes, since they can leave an uneven barrier and may require a longer spin or higher speed to separate fully. This is a separation-quality issue more than a direct hemolysis driver, but the wrong rotor invites the workarounds (longer, faster, repeated spins) that do raise free hemoglobin.
• Mechanical condition. A centrifuge that runs out of balance, vibrates, or builds heat on long runs is not performing as designed, and uneven or prolonged runs are the conditions that stress cells. Treat this as a maintenance and reliability question.

Why do my serum samples keep coming back hemolyzed?

If hemolyzed samples keep recurring, the spin step is usually not where the problem starts. The main drivers are upstream, at the draw and in transit:

• Line and catheter draws. In a large emergency-department study, the draw device was the strongest factor: blood taken through an IV-catheter hemolyzed about seven times as often as blood drawn with a butterfly needle. Small-gauge access and difficult venipuncture each added further risk (Wollowitz et al., 2013).
• Draw technique and needle gauge. Pulling blood too fast or through a needle that is too narrow subjects red cells to shear stress, the mechanical force that tears them.
• Transport, especially hospital pneumatic tube systems. Pneumatic tube delivery shakes and accelerates samples. The agitation, and the air-to-liquid mixing it causes ruptures cells before the tube reaches the lab.
• Time to spin. Long delays between draw and processing give cells time to degrade. Separation of serum or plasma from the cells should take place within 2 hours of collection to prevent erroneous test results unless conclusive evidence indicates that longer contact times do not contribute to result error (Becton Dickenson and Company; Greiner Bio-One).

These share a mechanism: shear stress, the tearing and bubble-collapse forces that mechanical handling imposes on fragile cells. The useful reframe for troubleshooting is a question. Is the centrifuge causing hemolysis, or is it revealing hemolysis that already happened? In most recurring cases it is the second. Spinning the sample is a step that makes existing free hemoglobin visible in the separated plasma. Chase the centrifuge and the rejection rate will not move, because the cause was a 25-gauge needle or a pneumatic tube run three floors up.

What centrifuge speed and time prevents hemolysis?

The most useful change a lab can make at the spin step is to specify force, not speed. Protocols written in RPM are ambiguous, because the force a sample actually feels depends on how far the tube sits from the center of rotation, the reference radius. Two centrifuges set to the same RPM can deliver very different forces.

The relationship is fixed:

RCF = 1.118 × 10⁻⁵ × r × (RPM)²

where r is the reference radius in centimeters and RCF is relative centrifugal force in units of gravity (×g).

A worked example shows why it matters. At 3,500 RPM with a reference radius of 15 cm, the sample feels about 2,055 ×g. The same 3,500 RPM with a reference radius of 10 cm delivers only about 1,370 ×g, a third less force, on a sample the protocol meant to treat identically. Specify 2,000 ×g instead of an RPM value, and both setups land on the same force.

Three practices prevent the centrifuge-related share of hemolysis:

• Specify RCF, not RPM. Match the reference radius so the delivered force is what the protocol intended, and validate spin parameters against CLSI guidance rather than carrying over a number from an old unit.
• Avoid re-spins. This is the single parameter most clearly tied to rising free hemoglobin.
• Confirm the rotor fits the tube. Use swing-out rotors for gel tubes, and verify speed and timer on a set schedule so the unit delivers what the display claims.

What is the best centrifuge for minimizing sample hemolysis?

The spin is the one stage of the preanalytical chain a lab fully controls, so the goal is to take the guesswork out of it. A centrifuge with a fixed, known relative centrifugal force and a rotor matched to the tube removes the two main spin-side variables: it delivers the force the protocol specifies, and it separates gel tubes cleanly without the longer or repeated spins that raise free hemoglobin. Reliable, balanced operation and dependable speed control keep the unit from becoming a variable in its own right.

Drucker Diagnostics builds clinical centrifuges around that predictability. See our Clinical Laboratory solutions to match a unit and rotor to your tube types and throughput.

References:

• Procedures for the Handling and Processing of Blood Specimens for Common Laboratory Tests (GP44-A4); and Handling, Transport, Processing, and Storage of Blood Specimens for Routine Laboratory Examinations (PRE04, 1st ed., 2023, which replaces GP44-A4). Wayne, PA.
• Becton, Dickinson and Company. BD Vacutainer Evacuated Blood Collection System: Instructions for Use. Franklin Lakes, NJ.
• Mancuso JE, Jayaraman A, Ristenpart WD. Centrifugation-induced release of ATP from red blood cells. PLoS One. 2018;13(9):e0203270. (University of California, Davis.)
• Heyer NJ, Derzon JH, Winges L, et al. Effectiveness of practices to reduce blood sample hemolysis in EDs: a laboratory medicine best practices systematic review and meta-analysis. Clin Biochem. 2012;45(13-14):1012-1032. doi:10.1016/j.clinbiochem.2012.08.002. (CDC Laboratory Medicine Best Practices review; Battelle, Seattle, WA.)
• Wollowitz A, et al. Use of butterfly needles to draw blood is independently associated with marked reduction in hemolysis compared to intravenous catheter. Acad Emerg Med. 2013;20(11):1151-1155. doi:10.1111/acem.12245. (Montefiore / Albert Einstein College of Medicine, NY.)
• Lippi G, Blanckaert N, Bonini P, et al. Haemolysis: an overview of the leading cause of unsuitable specimens in clinical laboratories. Clin Chem Lab Med. 2008;46(6):764-772.
• Getawa S, Aynalem M, Melku M, Adane T. Blood specimen rejection rate in clinical laboratory: a systematic review and meta-analysis. Pract Lab Med. 2023;33:e00303.
• Greiner Bio-One GmbH. VACUETTE® Evacuated Blood Collection System: Instructions for Use. Doc. 980200B Rev. 09. Kremsmünster, Austria; 2020.
• Wiegmann L, de Zélicourt DA, Speer O, Muller A, Goede JS, Seifert B, Kurtcuoglu V. Influence of standard laboratory procedures on measures of erythrocyte damage. Front Physiol. 2017;8:731. doi:10.3389/fphys.2017.00731.