Cell Fractionation

Cell fractionation is the process of disrupting a cell's plasma membrane under controlled conditions to release its internal contents, then separating those contents — organelles, membranes, macromolecular complexes — into distinct populations based on physical properties. The process has two conceptually separate stages. First, homogenization: mechanically shearing cells in a cold, isotonic (or slightly hypotonic) buffer so that the plasma membrane ruptures but organelles remain intact and functional. Second, centrifugation: spinning the resulting homogenate at controlled speeds to separate components by how fast they sediment through a fluid medium. Sedimentation rate depends on three things: the particle's size, its density relative to the surrounding medium, and the centrifugal force applied. Large, dense structures (nuclei, whole cells, cytoskeletal debris) sediment fastest; small, light structures (ribosomes, small vesicles) sediment slowest and require very high forces to pellet at all. Two centrifugation strategies exploit this differently. Differential centrifugation uses a series of increasing speeds on a homogeneous fluid, pelleting progressively smaller/lighter classes of organelles at each step — crude, fast, and good for a first pass. Density-gradient (equilibrium) centrifugation layers the homogenate atop a gradient of increasing density (commonly sucrose or Percoll) and spins long enough that each particle migrates to the point in the gradient matching its own buoyant density, regardless of size — much finer resolution, distinguishing organelles that differential spins would lump together. Critically, fractionation is only useful if fractions can be identified. Because you cannot simply look at a pellet and know what it is, each fraction is assayed using a marker — an enzyme or molecule known, from independent evidence, to reside uniquely (or predominantly) in one organelle. Co-purification of a marker enzyme's activity with a particular fraction is the operational proof that the fraction corresponds to that organelle. This logic — structure defined by function, and function assigned by biochemical assay rather than appearance — is what separates cell fractionation from simple microscopy, and it is why the technique became foundational to modern cell biology.

Step 1 — Choosing and preparing the tissue. Tissue (classically rat liver, chosen for its homogeneity and abundance of mitochondria) is minced and suspended in a cold buffer, typically isotonic sucrose (0.25 M) with a bit of buffer (e.g., Tris or HEPES) at pH ~7.4, plus sometimes EDTA to chelate divalent cations and inhibit degradative enzymes. Everything is kept at 0–4°C throughout. Cold temperature is essential: it slows the activity of hydrolytic enzymes (proteases, nucleases, lipases) that would otherwise degrade organelles and proteins once cellular compartmentalization is destroyed, and it reduces membrane fluidity, making organelles more mechanically robust during shearing.

Step 2 — Homogenization. The tissue is disrupted using a mechanical device — classically a Potter-Elvehjem homogenizer, a glass tube with a close-fitting Teflon pestle rotated and moved up and down, generating shear forces that rupture plasma membranes. Other methods include blenders (Waring blender, as used in the Hershey-Chase experiment — though there for a different purpose), sonication, or nitrogen cavitation. The goal is a homogenate: a mixture in which the plasma membrane is broken but internal organelle membranes (nuclear envelope, mitochondrial membranes, ER, Golgi, lysosomal membranes) remain intact and enclose their original contents. Osmotic balance matters enormously here — if the medium is too hypotonic, organelles like mitochondria and lysosomes will swell and lyse, releasing their enzymes into the surrounding fluid and contaminating every other fraction.

Step 3 — Differential centrifugation. The homogenate is spun in a series of steps of increasing centrifugal force (measured in units of gravity, ×g) and duration. After each spin, the pellet is removed and the supernatant is transferred to a new tube and spun again at higher speed. A typical scheme, following the pattern used by Claude and de Duve:

Each pellet is a crude fraction — not perfectly pure, since organelles of similar size/mass overlap somewhat between fractions — but progressively enriched. The nuclear fraction comes first because nuclei are by far the largest and densest organelle. Mitochondria (and lysosomes and peroxisomes, which are similar in size) come next. The lightest membrane fragments and ribosomes require the highest forces because their sedimentation rate is lowest.

Step 4 — Density-gradient centrifugation for finer separation. Because mitochondria, lysosomes, and peroxisomes are similar in size, differential centrifugation alone cannot cleanly separate them — they co-pellet. To resolve this, the crude organelle pellet is resuspended and layered on top of a density gradient: a tube pre-filled with sucrose solution whose concentration (and thus density) increases from top to bottom, prepared either as discrete steps or a continuous gradient. Upon centrifugation at high speed for a long time (hours), each particle migrates down the gradient until it reaches the depth at which the medium's density equals its own buoyant density — the point at which net force on the particle is zero. This is isopycnic (equal-density) centrifugation: separation is by density alone, independent of particle size or shape, given sufficient time to reach equilibrium. Peroxisomes and lysosomes, though similar in size to mitochondria, differ in density (their membrane-to-lumen composition differs), so equilibrium centrifugation separates them into distinct visible bands.

Step 5 — Assaying each fraction with markers. Once fractions are collected (by puncturing the tube bottom, or careful pipetting from the top down), each is tested for the presence of specific molecules:

The correlation between a fraction's position in the gradient and a marker's activity is the empirical proof of identity — this is how, for example, de Duve discovered the lysosome despite never having seen one in a micrograph beforehand.

Albert Claude — founding the method (1930s–1940s). Working at the Rockefeller Institute, Albert Claude set out to isolate the agent responsible for Rous sarcoma (a tumor virus) from chicken tissue, and in doing so developed the systematic use of differential centrifugation on cell homogenates. He homogenized cells and spun them at increasing speeds, discovering that he could reproducibly isolate distinct fractions — a heavy fraction containing nuclei, a "large granule" fraction (which we now call mitochondria) that showed strong respiratory (oxygen-consuming) enzyme activity, and a fraction of small particles rich in RNA that he called "microsomes" (later shown to be fragments of endoplasmic reticulum studded with ribosomes). Claude's key methodological insight was combining electron microscopy (to see what a fraction looked like) with biochemical assay (to see what it did) — establishing the marker-enzyme logic that defines the field. For this pioneering fractionation work, Claude shared the 1974 Nobel Prize in Physiology or Medicine with Christian de Duve and George Palade.

Christian de Duve — discovering the lysosome by pure biochemistry (1949–1955). De Duve was not looking for a new organelle; he was studying the enzyme glucose-6-phosphatase and its role in insulin action, and needed to measure another enzyme, acid phosphatase, as a control. He found something strange: when he assayed a crude liver homogenate immediately after preparation, acid phosphatase activity was low — but if he let the homogenate sit in the refrigerator for a few days, or subjected it to freeze-thaw or mild mechanical disruption, activity jumped dramatically. His interpretation, tested rigorously, was that the enzyme was latent — trapped inside an intact membrane-bound compartment that had to be physically ruptured before the enzyme could access its substrate in solution. Using differential centrifugation, he found acid phosphatase concentrated in a fraction distinct from mitochondria (identified by cytochrome oxidase activity, another marker), and that this new fraction's marker enzyme behaved as if enclosed in a membrane: activity was low unless the fraction was treated with detergent or osmotic shock to break the enclosing membrane. He named this novel organelle the lysosome ("lytic body") — a discovery made entirely through enzyme kinetics and fractionation, years before lysosomes were confirmed and visualized by electron microscopy. De Duve's group used the same logic shortly after to discover the peroxisome, initially detected as a fraction containing catalase and urate oxidase activity that banded at a different density than mitochondria or lysosomes in sucrose gradients — again, biochemistry led, structure followed. This body of work, done with George Palade's parallel electron-microscopic confirmations at Rockefeller, earned de Duve the 1974 Nobel Prize (shared with Claude and Palade) "for their discoveries concerning the structural and functional organization of the cell."

What these experiments proved, generally. They established three things that remain foundational: (1) organelles are not just static shapes seen in micrographs but discrete biochemical compartments with distinct, purifiable enzyme complements; (2) function can be discovered before structure is directly observed, through the logic of latency and marker co-purification; (3) careful control of osmotic and thermal conditions during homogenization is essential — sloppy preparation (e.g., excessive hypotonicity) ruptures organelles and destroys the very compartmentalization being studied, a pitfall de Duve had to overcome to correctly interpret his "latent enzyme" observations.

Example 1 — Interpreting an unknown gradient profile. A researcher homogenizes rat liver, isolates the crude "heavy" pellet from a 20,000 × g spin (expected to contain mitochondria, lysosomes, and peroxisomes), and runs it on a continuous sucrose density gradient (1.05–1.25 g/mL) at 100,000 × g for 3 hours. After fractionation, she measures three marker enzymes across the gradient (fraction 1 = top/light, fraction 12 = bottom/dense):

Reasoning: Succinate dehydrogenase is an inner mitochondrial membrane marker, so fraction 7 (density 1.18) is mitochondria-enriched. Acid phosphatase is the classic lysosomal marker, so fraction 3 (density 1.10, the lightest of the three) is the lysosome-enriched fraction. Catalase marks peroxisomes, so fraction 11 (density 1.23, the densest) is peroxisome-enriched. This ordering — lysosomes lightest, then mitochondria, then peroxisomes densest — matches the real physiological densities of these organelles in standard sucrose gradients, illustrating why isopycnic centrifugation, not differential centrifugation alone, was required to separate three organelles that co-pellet at 20,000 × g. If she had stopped at differential centrifugation, all three enzyme activities would have appeared mixed together in one crude pellet.

Example 2 — Diagnosing a homogenization error. A student homogenizes cells in distilled water (rather than isotonic sucrose) "to make lysis easier," then performs differential centrifugation and assays the low-speed supernatant (the fraction expected to be nearly organelle-free cytosol) for acid phosphatase activity. She finds strong activity there — a result inconsistent with acid phosphatase being a lysosomal (membrane-enclosed) enzyme. Reasoning: Distilled water is strongly hypotonic relative to the cytoplasm. Water rushes into organelles by osmosis, causing lysosomes (and other organelles) to swell and burst during homogenization, releasing their lumenal enzymes into the surrounding medium before any centrifugation occurs. Once acid phosphatase is free in solution, it behaves like a soluble cytosolic protein: it stays in the supernatant at every centrifugation step rather than pelleting with an intact lysosomal fraction. The fix is to homogenize in an isotonic medium (0.25 M sucrose is isosmotic with mammalian cytoplasm, ~300 mOsm) so organelles retain their native volume and membrane integrity, keeping the enzyme correctly latent until deliberately released for assay. This scenario is precisely the artifact de Duve had to rule out and control for when he first observed "latency" — distinguishing true membrane enclosure from simple freeze-thaw damage required careful, repeatable control experiments.

Christian de Duve was a Belgian cytologist and biochemist trained originally in medicine and endocrinology, whose early research on insulin's mechanism of action led him, almost by accident, into subcellular fractionation. Working in Louvain and collaborating closely with Albert Claude's group at the Rockefeller Institute in New York, de Duve brought a physician's insistence on rigorous, quantitative enzyme kinetics to a field that had previously relied mostly on visual description. His discovery of the lysosome — and, soon after, the peroxisome — exemplified a style of reasoning now central to cell biology: trust a biochemical anomaly (unexpectedly low, "latent" enzyme activity) enough to chase it methodically, using fractionation and marker logic, until it reveals an entirely new structure that no one had directly observed before. He coined the terms "lysosome," "peroxisome," and later "autophagy," and spent much of his later career studying lysosomal storage diseases — genetic disorders in which the very organelle he discovered fails to degrade its substrates properly, a direct clinical legacy of his fractionation work.

Cell fractionation is not a historical curiosity — it remains a working tool across biomedicine and biotechnology. In clinical diagnostics, fractionation-based enzyme assays help identify lysosomal storage diseases (e.g., Tay-Sachs, Gaucher disease), where a specific lysosomal enzyme is missing or non-functional; purifying the lysosomal fraction from patient cells and assaying enzyme activity directly is still part of definitive diagnosis. In cancer biology and cell signaling research, subcellular fractionation is used routinely to determine where a signaling protein localizes upon activation — for instance, showing that a transcription factor moves from cytosolic to nuclear fractions after a stimulus, evidence of nuclear translocation. In mitochondrial disease research, isolated mitochondrial fractions are assayed directly for electron transport chain complex activities to diagnose specific respiratory chain defects. In biotechnology and pharmaceutical development, fractionation underlies the purification of vaccines and viral vectors (density-gradient centrifugation separates intact virus particles from cellular debris and defective particles), and is a step in manufacturing biologics that require organelle-derived components. In basic research, fractionation feeds directly into modern organelle proteomics: purified fractions are subjected to mass spectrometry to catalog the complete protein complement of an organelle, refining and expanding the marker-enzyme approach with far greater resolution. The logic also underlies the biochemical purification of plasma membrane fractions used to study Membrane Proteins and to test predictions of The Fluid-Mosaic Model of Membranes, since isolating a pure membrane fraction was a prerequisite for ever characterizing lipid and protein composition of the membrane bilayer. Even CRISPR and gene-editing workflows often rely on fractionation-adjacent techniques (nuclear vs. cytoplasmic fractionation) to confirm where an edited protein or RNP complex localizes.

Ask the tutor: How would you redesign a fractionation protocol to separate two organelles that have nearly identical buoyant density?