The nephron begins with the glomerulus, which is composed of a capillary network lined by a thin layer of endothelial cells, a central region of mesangial cells with surrounding matrix material, and the visceral epithelial cells (podocytes) overlying the capillaries ( Figs. 2.7–2.10 ). The parietal layer of the Bowman capsule with its basement membrane encases the glomerulus. The Bowman space, or urinary space, is the cavity between the visceral and parietal epithelia. Although “renal corpuscle” is strictly the correct terminology to refer to the glomerulus and Bowman capsule, glomerulus is used throughout this chapter because of its common use. At the vascular pole, where the afferent and efferent arterioles enter and exit the glomerulus, the visceral epithelium is continuous with the parietal epithelium. The parietal epithelium transitions to the proximal tubule epithelium at or near the urinary pole. The average diameter of a glomerulus is approximately 200 μm in the human kidney and 120 μm in the rat kidney. However, the size and number of glomeruli vary significantly with age, gender, birth weight, and renal health. The average glomerular volume is 0.6 to 1 million μm ³ in rats ^, and 3 to 7 million μm ³ in humans, ^{, ,} although individual glomerular volume within single human kidneys can vary as much as eightfold. Rat juxtamedullary glomeruli are larger than superficial glomeruli; this is not the case in the human kidney.

Scanning electron micrograph of a rat glomerulus.

The glomerular tuft is encased by the Bowman capsule. Podocytes, with their interdigitating foot processes, cover the capillaries. The glomerular filtrate drains into the proximal tubule at the urinary pole.

From Sands JM, Verlander JW. Functional anatomy of the kidney. In: McQueen C, ed. Comprehensive Toxicology . 3rd ed. St. Louis: Elsevier; 2017.

Light micrograph of a normal glomerulus from a rat, demonstrating the four major cellular components: endothelial cell (E) , mesangial cell (M) , parietal epithelial cell (P) , and visceral epithelial cell or podocyte (V) .

The macula densa (MD) is in the thick ascending limb at the vascular pole.

Scanning electron micrograph of a cast of a glomerulus with its many capillary loops (CL) and adjacent renal vessels.

The afferent arteriole (A) takes its origin from an interlobular artery at lower left. The efferent arteriole (E) branches to form the peritubular capillary plexus (upper left).

Courtesy Waykin Nopanitaya, Ph.D.

Transmission electron micrograph of a normal rat glomerulus.

The capillary loops are lined by fenestrated endothelial cells (E), facing the capillary lumen (CL) . Mesangial cells (M) lie beneath the endothelial cells, among the capillary loops. Podocytes (P) and their extensive, interdigitating primary and secondary foot processes (arrows) cover the surface of the capillaries, facing the glomerular filtrate in the Bowman space (BS) . Parietal epithelial cells (PEC) line the Bowman capsule facing the BS.

The main function of the glomerulus is filtration of the plasma. The layers of the glomerular capillary wall, the fenestrated capillary endothelium, the glomerular basement membrane (GBM), and the filtration slit diaphragm between the foot processes of the visceral epithelial cells form the filtration barrier between the blood and the urinary space ( Fig. 2.11 ). To cross the capillary wall, a molecule must pass sequentially through the fenestrated endothelium, GBM, and filtration slit diaphragm. Although the glomerular capillary wall allows passage of small molecules, the prevailing view is that it normally restricts the passage of cells and larger molecules, such as albumin, due to its size- and charge-selective properties.

Transmission electron micrograph of normal rat glomerular capillary wall fixed in a 1% glutaraldehyde solution containing tannic acid.

Note the relationship among the three layers of the glomerular basement membrane and the presence of the pedicels (P) embedded in the lamina rara externa (arrowhead) . The filtration slit diaphragm with the central dense spot (thin arrow) is especially evident between the individual pedicels. The fenestrated endothelial lining of the capillary loop is shown below the basement membrane. A portion of an erythrocyte is in the extreme lower right corner. BS, Bowman space; CL, capillary lumen.

The glomerular capillaries are lined by a thin fenestrated endothelium ( Figs. 2.11 and 2.12 ). These endothelial cells form the initial barrier to passage of blood constituents from the capillary lumen to the Bowman space. Under normal conditions, the formed elements of the blood including erythrocytes, leukocytes, and platelets do not gain access to the subendothelial space.

Scanning electron micrograph of a glomerular capillary from the kidney of a normal rat.

Numerous endothelial pores, or fenestrae, are present in the endothelial cells lining the capillary lumen. The ridgelike structures are localized thickenings of the endothelial cells. Interdigitating foot processes of the podocytes cover the urinary side of the capillaries.

The endothelial cell nucleus lies adjacent to the mesangium, with the remainder of the cell irregularly attenuated around the capillary lumen (see Fig. 2.10 ). The endothelium contains pores or fenestrae that range from 70 to 100 nm in diameter in humans (see Figs. 2.11 and 2.12 ). Nonfenestrated, ridgelike structures termed “cytofolds” are found near the cell borders. An extensive network of intermediate filaments and microtubules is present in the endothelial cells, and microfilaments surround the fenestrations. Most studies indicate adult glomerular endothelial cells lack diaphragms across the fenestrae, whereas diaphragmed fenestrae are present in the embryo, where they may compensate for the functional immaturity of the embryonic glomerular filtration barrier. The glomerular endothelium is covered by a glycocalyx layer, the visualization of which requires special methods such as electron microscopy with cationic dyes or lipid particles. The glycocalyx also fills the endothelial fenestrae forming “sieve plugs,” the exact function of which is unknown. The glycocalyx consists of membrane-bound proteoglycans (syndecan and glypican) with attached glycosaminoglycans (GAGs), secreted glycoproteins (perlecan and versican), and secreted GAGs (hyaluronan), which provide a negative charge.

Classic ultrastructural studies demonstrated that endogenous albumin is largely confined to the glomerular capillary lumen and does not pass through the endothelium. In recent years, more studies have addressed the potential role of the glomerular endothelium, particularly its glycocalyx, in filtration. Studies in rats showed that eluting molecular components of the glomerular endothelial glycocalyx with hypertonic sodium chloride induced a 12-fold increase in proteinuria. Injection of hyaluronidase, a hyaluronan-degrading enzyme, in mice led to disruption of the glomerular endothelial glycocalyx and leakage of albumin across the endothelium. Using isolated human and rodent glomeruli, enzymatic disruption of the glomerular endothelial glycocalyx resulted in increased glomerular albumin permeability. Thus experimental evidence supports that the glomerular endothelial glycocalyx is an important component of the filtration barrier.

Signaling between glomerular cells is critical for the development and maintenance of the filtration barrier. The surfaces of glomerular endothelial cells express receptors for the vascular endothelial growth factor (VEGF) family. VEGF is synthesized by podocytes (glomerular visceral epithelial cells) and is an important regulator of microvascular permeability. ^, VEGF increases endothelial cell permeability and induces the formation of endothelial fenestrations. ^, VEGF-A is the best characterized podocyte growth factor, and its principal receptor is VEGFR2, expressed on endothelial cells. Podocyte-specific alterations of VEGF-A have demonstrated that it is required for normal differentiation of glomerular endothelial cells. ^, Moreover, drug inhibition of VEGF-A in patients or podocyte-specific deletion of VEGF-A in adult mice results in severe glomerular endothelial injury and thrombotic microangiopathy. Thus VEGF produced by podocytes plays a critical role in the differentiation and maintenance of glomerular endothelial cells and is an important regulator of endothelial cell permeability.

Several other cell–cell communication pathways exist between glomerular cells. For example, angiopoietin–TIE signaling regulates endothelial homeostasis in an intricate manner. Angiopoietin-1 (ANGPT1) produced in podocytes binds to endothelial-expressed tyrosine kinase receptor TIE2, the phosphorylation of which promotes endothelial survival. In contrast, angiopoietin-2 secreted by endothelial cells is an antagonist of ANGPT1-mediated TIE2 activation in endothelial cells.

By transmission electron microscopy, the GBM is composed of a central dense layer, the lamina densa, and two thinner, more electron-lucent layers, the lamina rara externa and the lamina rara interna (see Fig. 2.11 ). The latter two layers measure approximately 20 to 40 nm in thickness. Although in the rat the width of the GBM has been found to be 132 nm, the width of the human GBM has consistently been reported to be more than 300 nm ^, with a slightly thicker basement membrane in men (373 nm) than women (326 nm). Compared with other basement membranes, the GBM is thicker, likely at least in part from fusion of endothelial and epithelial basement membranes during development. Mass spectrometry–based proteomic analysis has revealed at least 212 proteins in the normal human glomerular extracellular matrix; however, like other basement membranes in the body, the GBM is composed primarily of type IV collagen, laminin, nidogen (entactin), and heparan sulfate proteoglycans (HSPGs).

Type IV collagen consists of six chains, α1(IV) through α6(IV). Three α(ΙV) chains self-associate intracellularly to form triple helical molecules called protomers. Three types of promoters are formed: α1α2α1, α3α4α5, and α5α6α5. Upon secretion into the extracellular space, the protomers self-associate via their amino- and carboxy-terminal domains to form polymerized networks. Three sets of collagen IV networks form: α1α2α1(IV)-α1α2α1(IV), α3α4α5(IV)-α3α4α5(IV), and α1α2α1(IV)-α5α6α5(IV). The networks undergo specific extracellular modifications to form an elaborate scaffold that tethers other molecules, serves as a cell-signaling interface, and provides support for adjacent cells. Halogens facilitate the assembly of the collagen IV scaffold. For example, extracellular chloride ions activate a molecular switch that enables individual promoter carboxy domains to oligomerize, and ionic bromide is essential for enzymatic cross-linking to stabilize the protomer–protomer connections. The α3α4α5(IV)-α3α4α5(IV) network predominates in the GBM, whereas the α1α2α1(IV)-α5α6α5(IV) network is in the Bowman capsule. Whereas α1α2α1(IV) protomers are synthesized from both endothelial cells and podocytes, α3α4α5(IV) protomers are secreted only by podocytes. Mutations in the genes encoding α3, α4, and α5(IV) chains cause Alport syndrome, and autoantibodies against the carboxy terminal a3(IV) chain are responsible for anti-GBM disease.

Laminins (LMs) are large heterotrimeric glycoproteins composed of three chains: α, β, and γ. The major laminin in the adult GBM is LM-521 (containing the α5, β2, and γ1 chains). Both glomerular endothelial cells and podocytes synthesize laminin α5 and β2. Mutations of laminin β2 result in a congenital nephrotic syndrome called Pierson syndrome in humans.

Nidogens, also known as entactins, are glycoproteins. Nidogen-1 binds to both collagen IV and laminin but does not appear essential for GBM formation. HSPGs consist of a core protein linked to sulfated GAG side chains. Agrin, perlecan, and type XVIII collagen are HSPGs found in the GBM. Agrin is the major HSPG in the GBM, whereas perlecan and type XVIII collagen are found mainly in the mesangium.

Subdiffraction resolution stochastic optical reconstruction microscopy has provided a precise view of the nanoscale organization of these molecular networks within the GBM. The α3α4α5(IV) network localizes to the center of the GBM, whereas the α1α2α1(IV) network maps near the endothelial side of the GBM. Laminin-521 is situated in two layers near the endothelial and podocyte sides of the GBM and in the central portion of the GBM. Agrin localizes in two layers along the endothelial and podocyte surfaces of the GBM, with more detected near the podocytes.

The contribution of the GBM to the glomerular filtration barrier has been studied for decades. ^, Ultrastructural tracer studies provided evidence to suggest that the GBM constitutes both a size-selective and a charge-selective barrier. Additional studies revealed a lattice of anionic sites with a spacing between them of approximately 60 nm ( Fig. 2.13 ) throughout the lamina rara interna and lamina rara externa. ^, The anionic sites in the GBM consist of heparan sulfate GAG side chains of the proteoglycans rich in heparan sulfate. ^, Removal of the heparan sulfate side chains by enzymatic digestion resulted in an increase in the in vitro permeability of the GBM to ferritin and to bovine serum albumin, suggesting that HSPGs play a role in establishing the permeability properties of the GBM to plasma proteins (see Fig. 2.13 ). However, in vivo studies have addressed the role of proteoglycans and charge selectivity in the GBM. Overexpression of heparinase in transgenic mice led to a fivefold reduction in GAG-associated sites in the GBM but no proteinuria. Moreover, podocyte-specific deletion of agrin alone or in combination with deletion of perlecan heparan sulfate side chains in mice resulted in a dramatic reduction in GBM anionic sites but did not alter the filtration barrier to albumin or a negatively charged tracer. ^, Thus more recent data suggest that the role of GBM anionic charge, at least that contributed by proteoglycans, is minimal in the function of the glomerular filtration barrier.

Transmission electron micrographs of the glomerular filtration barrier in normal rats perfused with native anionic ferritin (A) or cationic ferritin (C) and in rats treated with heparitinase before perfusion with anionic (B) or cationic ferritin (D).

In normal animals, anionic ferritin is present in the capillary (Cap) but does not enter the glomerular basement membrane (GBM), as shown in (A). In contrast, cationic ferritin binds to the negatively charged sites in the lamina rara interna (LRI) and lamina rara externa (LRE) of the GBM (see C). After treatment with heparitinase, both anionic (B) and cationic (D) ferritin penetrate the GBM, but there is no labeling of negatively charged sites by cationic ferritin. En, Endothelial fenestrae; Fp, foot processes; LD, lamina densa; US, urinary space.

Modified from Kanwar YS. Biophysiology of glomerular filtration and proteinuria. Lab Invest. 1984;51:7–21.

Nevertheless, a variety of genetic findings in humans and studies in mice indicate that an intact GBM serves a barrier function to protein permeability. The absence of an intact α3α4α5(IV) network in the GBM of Alport syndrome eventually results in proteinuria. In humans and animal models of Alport syndrome, there is a compensatory increase in the α1α2α1(IV) network, laminin α5 chain, and ectopic laminin isoforms (α1, α2, and β1 chains) in the defective GBM. ^, These secondary changes alter cell–matrix signaling, are accompanied by characteristic splitting and “basket-weave” lamellation of the GBM, and produce proteinuria.

Strong evidence for a specific role of the GBM in the filtration barrier is the presence of laminin β2 mutations in humans or mice resulting in massive proteinuria. ^, Laminin-β2–deficient ( Lamb 2 ^–/– ) mice develop severe proteinuria and ectopic laminin chains (α1, α2, α3, β3, and γ2) that accumulate in the GBM, but this ectopic deposition fails to compensate for the absence of laminin β2. Importantly, the albuminuria in the mice precedes podocyte foot process effacement and filtration slit diaphragm abnormalities, indicating the GBM has an essential role in the filtration barrier. Remarkably, injection of recombinant human LM-521 accumulates in the correct orientation in the GBM and delays the onset of proteinuria in Lamb 2 ^–/– mice, which lack LM-521.

Podocytes (visceral epithelial cells) are the largest cells in the glomerulus and are positioned on the outside of the glomerular capillary wall (see Figs. 2.7 , 2.10–2.12 , and 2.14 ). Mature podocytes are terminally differentiated and generally do not replicate. They have a prominent cell body containing nuclei, endoplasmic reticulum, Golgi apparatus, and an endocytic–lysosomal system. The cell bodies give rise to long cytoplasmic primary processes that branch into secondary and tertiary processes, surround the capillaries, and finally divide into foot processes. The foot processes come into direct contact with the lamina rara externa of the GBM (see Figs. 2.11 and 2.13 ). By scanning electron microscopy (SEM), it is apparent that adjacent foot processes are derived from different podocytes (see Fig. 2.14 ). The gap between adjacent foot processes is bridged by a thin structure called the “filtration slit diaphragm.” Advanced techniques, including serial block-face SEM (SBF-SEM) and focused ion beam SEM (FIB-SEM), show that foot processes emerge directly from the podocyte cell body and elongated cytoplasmic processes. ^, These studies reveal tortuous ridgelike prominences along the basal surface of the cell body and cytoplasmic processes, from which the proximal portions of the foot processes emerge.

Scanning electron micrograph of a glomerulus from the kidney of a normal rat.

The visceral epithelial cells, or podocytes (P), extend multiple processes outward from the main cell body to wrap around individual capillary loops. Immediately adjacent pedicels, or foot processes, arise from different podocytes.

A series of studies using three-dimensional (3D) electron microscopic reconstruction and SBF-SEM have supported the existence of a subpodocyte space (SPS) under the podocyte cell body and a narrow interpodocyte space, which interconnects the SPS with the peripheral Bowman space. ^, Whether these spaces act as a resistance pathway across the filtration barrier remains to be determined.

Podocytes have an elaborate cytoskeleton that underlies their shape, stability, adhesion, and response to stress. Large numbers of microtubules and intermediate filaments (vimentin) are present in the cell body and primary processes, whereas actin filaments are especially abundant in the foot processes. Ultrastructural studies have demonstrated two distinct actin filament networks in foot processes of rat podocytes. ^, “Actin bundles,” containing α-actinin and synaptopodin, extend along the longitudinal axis of the foot processes above the level of the slit diaphragm. The cortical actin network, containing cortactin, lies between the actin bundle and plasma membrane. In glomerular diseases associated with proteinuria, the podocyte cytoskeleton is disrupted, slit diaphragms are lost, and the interdigitating foot processes are replaced by broad regions of podocyte processes covering the GBM. This “foot process effacement” is often accompanied by aggregated filaments appearing as a cytoplasmic mat juxtaposed to the GBM.

Actin fibers are composed of bundles of actin filaments. Both contractile and noncontractile actin fibers are present in most cells, and the former (actin stress fibers) are characterized by periodic alternating bands of α-actinin and myosin. Studies using super-resolution microscopic methods have revealed a detailed model of the podocyte actin cytoskeleton in both mice and humans. Actin fibers in the center of the foot process contain α-actinin and synaptopodin but lack myosin IIA, whereas actin fibers in the podocyte cell body and primary processes contain myosin IIA but lack synaptopodin. These findings suggest the actin fibers in the foot process are noncontractile, whereas the actin fibers in the cell body and primary processes are contractile. In podocyte injury models with foot process effacement and proteinuria, myosin IIA translocates to cytoplasm adjacent to the GBM, forming sarcomere-like structures with alternating synaptopodin and α-actinin staining. Thus podocytes contain distinct actin filament networks that appear to provide tensional integrity (tensegrity) and can redistribute in response to injury.

Foot processes contain two structures, focal adhesions (FAs) and filtration slit diaphragms (SDs), that interact with and control the actin cytoskeleton. FAs anchor the base of the foot processes to the GBM. They consist of transmembrane protein complexes through which the actin cytoskeleton is regulated by extracellular signaling. Cell adhesions rich in integrins and their interacting proteins are known as the “integrin adhesome.” FAs are a form of integrin adhesome and, at the foot process–GBM interface, consist of α3β1 integrin and various adaptor proteins, kinases, phosphatases, and guanosine triphosphatases (GTPases). The α3β1 integrin interconnects laminin in the GBM with the talin, paxillin, and vinculin adaptor cytoplasmic complex, which link to the actin cytoskeleton. Mutations of the integrin α3 subunit in humans are associated with massive proteinuria. Podocyte adhesion to the GBM is supported by the interaction of integrin α3β1 with the tetraspanin protein CD151, the absence of which leads to severe proteinuria. FA kinase and integrin-linked kinase localize to FAs and mediate signaling with the actin cytoskeleton.

The Rho family of small GTPases including RhoA, Rac1, and Cdc42 regulate actin cytoskeleton dynamics. Activation of podocyte RhoA and Rac1 in transgenic mice leads to proteinuria and podocyte foot process effacement. In contrast, podocyte deletion of C4dc42 results in proteinuria and foot process effacement. ^, Rho GTPases cycle between an active GTP-bound form and an inactive guanosine diphosphate (GDP)-bound form. Rho GTPases are inactivated by GTPase-activating proteins (GAPs), which increase GTP hydrolysis, or by guanine nucleotide dissociation inhibitors (GDIs), which sequester their inactive GDP-bound form in the cytoplasm. Mutations in the genes that encode Arhgap24, a GAP, and Arhgdia, a GDI, result in Rac1 activation and proteinuria in humans. ^, Moreover, mutations in Kank2 (kidney ankyrin repeat-containing protein), an Arhgdia -interacting protein that localizes to FAs, lead to RhoA activation and proteinuria. Dynamin is a large GTPase that has a role in clathrin-mediated endocytosis but also directly binds actin filaments and promotes actin polymerization. Dynamin regulates FA maturation in podocytes in vitro, and its conditional deletion in mouse podocytes results in foot process effacement and severe proteinuria. ^, These studies suggest that dynamin serves as a molecular link between endocytosis and actin remodeling in podocytes. Thus an intricate physiologic balance of the various GTPases is required for normal podocyte homeostasis.

The filtration SD is the second structure that controls the actin cytoskeleton. It appears as a thin line on electron microscopy (see Fig. 2.11 ) and bridges the 30–40 nm space (called the filtration slit) between adjacent foot processes. A central dot within the SD may occasionally be seen on ultrastructural cross-sections, and it appears as a continuous central filament on sections parallel to the plane of the GBM ( Fig. 2.15 ). On the basis of these observations, Rodewald and Karnovsky proposed a porous zipper-like model for the SD. In this model, there are regularly spaced cross-bridges that extend from the membranes of two adjacent foot processes to a linear central filament that runs equidistant and parallel to the cell membranes. The cross-bridge structures measuring 7 × 14 nm are separated by pores measuring 4 × 40 nm. Advanced microscopy techniques have provided alternative models and insights into the SD structure. On the basis of a freeze-etching replica ultrastructural method, it was proposed that the SD has a sheetlike rather than a zipper-like substructure. Electron microscope tomography showed the SD to consist of a network of winding cross strands, 30–35 nm in length, which merge centrally into a longitudinal density. Although this study generally concurred with the zipper-like model, the pores surrounding the strands appeared more irregular than originally proposed. In contrast, an investigation using enhanced SEM revealed variable-shape pores in the center of the SD and no central filament. This finding is more consistent with the SD as a heteroporous structure rather than the zipper-like model. High-resolution helium-ion SEM studies demonstrate the SD with cross-bridging filaments and surrounding pores forming a ladderlike structure in the middle of the filtration slit, also without a distinct central midline, thus generally supporting the heteroporous model. ^, The complexity of the SD is further illustrated by cryo-EM tomographic studies showing distinct cross-bridging strands are composed of different molecules. Bridging shorter strands in the lower part of the SD closest to the GBM consist of the nephrin-related protein, Neph1, whereas longer strands in the top part of the SD toward the apical side contain nephrin. This study supports the existence of a layered bipartite molecular assembly within the SD.

Electron micrograph showing the epithelial foot processes of normal rat glomerulus preserved in a 1% glutaraldehyde solution containing tannic acid.

In several areas, the slit diaphragm has been sectioned parallel to the plane of the basement membrane, revealing a highly organized substructure. The thin central filament corresponding to the central dot observed on cross-section (see Fig. 2.11 ) is indicated by the arrows.

Our understanding of the podocyte role and its SD in the filtration barrier was accelerated with identification of the protein nephrin, encoded by NPHS1, the gene mutated in congenital nephrotic syndrome of the Finnish type. Nephrin normally localizes to the SD, and its absence in the human congenital syndrome or in transgenic mice leads to loss of the SD, foot process effacement, and massive proteinuria. The SD area or domain of the podocyte includes the SD itself and the adjacent foot process membrane and cytoplasm. An expanding number of proteins localize to the SD domain, where they interact with nephrin and other partners, forming a multiprotein complex. Mutations in more than 30 genes, many of which localize to the SD domain and the podocyte actin cytoskeleton, cause human nephrotic syndrome. For example, mutations or deficiencies of genes encoding SD domain proteins, such as podocin, CD2-associated protein, phospholipase Cε1, and transient receptor potential cation channel type 6, result in SD loss, foot process effacement, and proteinuria. Thus there is convincing genetic evidence for the essential role of the SD, likely as a size-selective element, in the filtration barrier.

In addition to functioning as a critical structural barrier in filtration, the SD also functions as a signaling hub to regulate actin dynamics. Although the signaling pathways are incompletely understood, nephrin plays a central role. For example, phosphorylation of tyrosine residues within the intracellular domain of nephrin by Fyn kinase results in the recruitment of actin adaptor proteins such as Nck proteins (Nck1 and Nck2), which, in turn, induce actin polymerization. ^, Moreover, Nck protein binding to Fyn promotes increased phosphorylation of nephrin. Downstream of its interaction with nephrin, Nck directly binds to and activates the neuronal Wiskott-Aldrich syndrome protein (N-WASP), an actin nucleation protein. N-WASP binds and activates the ubiquitously expressed Arp2/3 multiprotein complex, which induces actin polymerization. ^, The importance of this signaling pathway is highlighted by studies showing that intact nephrin phosphorylation and the presence of podocyte Nck and N-WASP proteins are required for an intact filtration barrier of foot processes and stabilization of foot processes. There is also increasing evidence that the phosphorylation state of nephrin plays a role in its endocytic trafficking within the podocyte and is important for turnover and maintenance of the SD.

The mesangial cells and their surrounding matrix constitute the mesangium, which provides a scaffold for the surrounding glomerular capillaries. ^{, ,} The mesangium is separated from the capillary lumens by the endothelium and is surrounded by the GBM between capillary loops (see Figs. 2.8 and 2.10 ). Thus the mesangium directly abuts both the endothelium and the GBM. The points where the GBM no longer encircles the capillary and starts to surround the mesangium are called the “mesangial angles.” Three-dimensional reconstruction studies reveal continuity of the entire mesangium as a continuous arborizing structure within the glomerulus. Mesangial cells are located within the central axial region of the mesangium and are irregular in shape with a dense nucleus. They have elongated cytoplasmic processes that extend toward the endothelium and the adjacent GBM (paramesangial GBM). At the endothelial interface, the fingerlike mesangial cell processes may extend a short distance into the space between the endothelium and GBM. In certain forms of glomerular injury, the mesangial processes may insinuate between the endothelium and GBM for a noticeable distance along the peripheral capillary wall (mesangial interposition). In addition to the usual complement of organelles, mesangial cells possess an extensive array of microfilaments containing actin, myosin, and α-actinin. The mesangial processes, containing bundles of microfilaments, appear to bridge the gap in the GBM encircling the capillary, adhere to endothelial cells, and interconnect opposing mesangial angles of the GBM. This cell–matrix interconnection is believed to prevent capillary wall distention secondary to elevation of the intracapillary hydraulic pressure.

Several studies have elucidated molecules that mediate the interactions between mesangial cells and other glomerular cells and also the GBM. Afadin, an F-actin binding protein, localizes to cell contacts between mesangial and endothelial cells, also colocalizes with β-catenin, and may play a role in mesangial cell migration. Integrin α3β1 and Lu/BCAM are mesangial receptors that mediate adhesion of mesangial cells to the laminin α5 chain in the GBM. An actin cross-linking protein, EPLIN, is highly expressed in mesangial cell processes at the mesangial angles, where they attach to the GBM. Nephronectin, a protein within the GBM, binds to its receptor a8b1 integrin, produced by mesangial cells to form a GBM–mesangial adhesion at the lateral base (near the mesangial angles) of the capillary loops.

The mesangium is continuous with the extraglomerular mesangium, a component of the juxtaglomerular apparatus (JGA). The intraglomerular and extraglomerular cells are similar, and gap junctions exist between them. Cells of renin lineage within the extraglomerular mesangium have been shown to migrate and repopulate the mesangium after glomerular injury.

As proposed by Schlondorff, the mesangial cell may have some specialized features of pericytes and possesses many of the functional properties of smooth muscle cells. In addition to providing structural support for the glomerular capillary loops, the mesangial cell has contractile properties and is thought to play a role in the regulation of glomerular filtration. The local generation of autacoids, such as prostaglandin E ₂ , by the mesangial cell may provide a counterregulatory mechanism to oppose the effect of vasoconstrictors.

Mesangial cells exhibit phagocytic properties and participate in the clearance of macromolecules from the mesangium, ^, as evidenced by the uptake of tracers such as ferritin, colloidal carbon, and aggregated proteins. Mesangial cells are also involved in the generation and metabolism of the extracellular mesangial matrix. ^, Because of both their distinct anatomic localization and their production of various vasoactive substances (e.g., nitric oxide), growth factors (e.g., VEGF, platelet-derived growth factor [PDGF], transforming growth factor [TGF]), and cytokines and chemokines (interleukins, chemokine [C-X-C motif] ligand 1, chemokine [C-C motif] ligand [CCLs]), mesangial cells are also perfectly suited to mediate an extensive crosstalk to both endothelial cells and podocytes to control and maintain glomerular function. The PDGF-B isoform, the main ligand for the receptor PDGFR-β, is a potent mitogen for mesangial cell proliferation, and genetic deletion of PDGF-B and PDGFR-β results in an absence of mesangial cells and mesangium. As such, the mesangial cells also importantly contribute to a number of glomerular diseases including IgA nephropathy and diabetic nephropathy.

The mesangial cell is surrounded by a matrix that is similar but not identical to the GBM; the mesangial matrix is more coarsely fibrillar and slightly less electron dense. The presence of abundant thin microfibrils, best observed with tannic acid staining, likely explains the fibrillary character of the mesangial matrix. Fibrillin-1 is the major protein of the microfibrils, but other associated proteins include microfibril-associated glycoproteins 1 and 2 and latent TGF-binding protein-1. ^, Fibrillin-1 and α8 integrin colocalize in the mesangium and appear to interact to regulate mesangial adhesion.

The mesangial matrix also contains fibronectin, type IV collagen α1 and α2 chains (not type IV α3, α4, or α5 chains, which are present in the GBM), type V collagen, various laminin isoforms (not laminin-521, which is present in the GBM), and the proteoglycan perlecan (not the proteoglycan agrin, which is present in the GBM). For example, laminin α1 is present in the mesangial matrix (not the GBM), and studies suggest it regulates mesangial cell homeostasis and matrix deposition by inhibiting TGF-β/Smad pathway signaling. Several cell surface receptors of the β-integrin family have been identified on the mesangial cells, including α1β1, α3β1, and the fibronectin receptor, α5β1. These integrins mediate attachment of the mesangial cells to specific molecules in the extracellular mesangial matrix and link the matrix to the cytoskeleton. The attachment to the mesangial matrix is important for cell anchorage, contraction, and migration; ligand–integrin binding also serves as a signal transduction mechanism that regulates the production of extracellular matrix, as well as the synthesis of various vasoactive mediators, growth factors, and cytokines. ^,

The parietal epithelium, which lines the inner aspect of the Bowman capsule, consists of flat squamous-like cells known as parietal epithelial cells (PECs) (see Fig. 2.10 ). At the urinary pole, there is an abrupt transition from the PECs to the taller cuboidal cells of the proximal tubule, which has a well-developed brush border ( Fig. 2.16 ). The PECs are 0.1 to 0.3 μm in height, except at the nucleus, where they increase to 2.0 to 3.5 μm. Each cell has a long cilium, and organelles are generally sparse but include small mitochondria, numerous vesicles of 40 to 90 nm in diameter, and the Golgi apparatus. Large vacuoles and multivesicular bodies are rare. PECs express Pax-2 and claudin-1. The thickness of the basement membrane of the Bowman capsule varies from 1200 to 1500 nm. The basement membrane often has a lamellated appearance and increases in thickness with disease processes. At both the vascular pole and the urinary pole, the thickness of the Bowman capsule decreases markedly. In contrast to the GBM, the basement membrane of the capsule contains the α6 chain of type IV collagen, which is part of the α1α2α1 (IV)-α5α6α5(IV) protomer network.

Scanning electron micrograph showing the surface of the parietal epithelial cells adjacent to the early proximal tubule (PT) at the urinary pole.

Parietal epithelial cells have single cilia, and their lateral cell margins are accentuated by short microvilli (arrowheads) .

PECs function as a permeability barrier for the urinary filtrate. In experimental glomerulonephritis, this barrier is compromised and macromolecules can leak into the space between the PECs and the basement membrane of the Bowman capsule and subsequently into the periglomerular space. Several investigations suggest different populations of PECs exist. Cells located at the glomerular vascular pole interposed between the PECs and podocytes have been called “peripolar cells.” By electron microscopy, these cells have prominent cytoplasmic granules, and display an immunophenotype between PECs and podocytes and are currently called “transitional cells.” ^, Their function is unknown. Other cells lining the Bowman capsule near the vascular pole expressing podocyte markers and forming interdigitating foot processes are called “parietal podocytes” (or “ectopic podocytes”). ^, In glomerular disease, PECs may transform into cuboidal cells with enlarged nuclei and express CD44. These “activated PECs” demonstrate proliferation, migration, and matrix deposition and play a role in diseases such as focal segmental glomerulosclerosis.

Several studies have addressed the role of PECs as possible progenitor cells to renew podocytes. These various investigations using different experimental mouse models of podocyte depletion and genetic labeling methods have shown that PECs may serve as podocyte progenitor cells, transdifferentiating into podocytes, which repopulate the glomerular tuft. ^, In some studies of experimental glomerular injury, a subpopulation of podocytes actually migrates to the Bowman capsule and expresses PEC markers. An understanding of these bidirectional differentiation pathways involving PECs and podocytes awaits further investigations.

The juxtaglomerular granular cells are located primarily in the walls of the afferent and, less commonly, the efferent arterioles. They exhibit features of both smooth muscle cells and secretory epithelial cells and therefore have been called epithelioid or myoepithelial cells. They contain myofilaments in the cytoplasm, a well-developed endoplasmic reticulum, and small “protogranules” with a crystalline substructure in the Golgi complex. ^, The signature feature of juxtaglomerular cells is the numerous electron-dense, membrane-bound granules of variable size and shape ( Fig. 2.17 ), which contain the aspartyl protease renin. ^, In addition to renin granules, lipofuscin-like granules are common in juxtaglomerular cells in the human kidney, as well as in extraglomerular mesangial cells. ^,

Transmission electron micrograph of juxtaglomerular apparatus from a rabbit kidney, illustrating macula densa (MD) , extraglomerular mesangium (EM) , and a portion of an arteriole (right) , containing numerous electron-dense granules.

Macula densa cells are significantly taller and narrower than the adjacent thick ascending limb (TAL) cells.

In addition to renin, the juxtaglomerular granular cells express angiotensin II, which localizes in the same granules as renin and has highest activity in the afferent arteriole. Like lysosomes, renin-containing granules have an acid pH and contain lysosomal enzymes, including acid phosphatases and cathepsin B, and have the capacity to take up and degrade internalized material. ^{, ,} During kidney development, renin expression is present in the intrarenal arteries, but by adulthood in normal conditions, renin granules are largely found only in juxtaglomerular granular cells in the distal afferent arteriole. Nonetheless, in adults, renin expression may again extend into more proximal arterial portions in some conditions, such as extravascular volume depletion, hypotension, and hemorrhage.

Located between the afferent and efferent arterioles in close contact with the macula densa (see Fig. 2.17 ), the extraglomerular mesangium is continuous with the intraglomerular mesangium and composed of cells that are similar in ultrastructure to the mesangial cells. ^, The extraglomerular mesangial cells possess long, thin cytoplasmic processes separated by basement membrane material. Although not typical, extraglomerular mesangial cells occasionally contain renin granules. The extraglomerular mesangial cells are in contact with the afferent and efferent arterioles and the macula densa, and gap junctions are commonly observed between the various cells of the vascular portion of the JGA. ^, Gap junctions, formed at least in part from connexin 40, exist between extraglomerular and intraglomerular mesangial cells, enabling signaling to be conveyed from the macula densa through the extraglomerular mesangium to the glomerulus. ^, Moreover, there is evidence that altered gap junction structure and function may eliminate the tubuloglomerular feedback response. ^,

The macula densa is a specialized region in the TAL adjacent to the hilum of the parent glomerulus (see Figs. 2.8 and 2.17 ). Macula densa cells are morphologically distinct from the surrounding cells of the TAL. They are columnar cells with large, apically placed nuclei, although there are considerable species differences in the height of macula densa cells. Compared with TAL cells, macula densa cells have relatively little cytoplasm, few basolateral plasma membrane infoldings, and lower mitochondrial density; mitochondria are small and either scattered (rat) or basal to the nucleus (human) and rarely enclosed within basolateral plasma membrane infoldings. The Golgi apparatus is lateral to and beneath the cell nucleus and other cell organelles including lysosomes, autophagic vacuoles, ribosomes, and smooth and rough endoplasmic reticulum and is also located principally beneath the cell nucleus. Basal cytoplasmic extensions contact the vascular elements, and at these points, the macula densa basement membrane is fused with basement membrane of the vascular elements. ^, Macula densa cells lack the lateral cell processes and interdigitations that are characteristic of the TAL, and the width of the lateral intercellular spaces varies with the physiologic state of the animal.

Cells of the PCT are structurally complex. ^{, ,} Large primary ridges extend laterally from the apical to the basal surfaces of the cells. Large lateral processes, often containing mitochondria, extend outward from the primary ridges and interdigitate with similar processes in adjacent cells ( Fig. 2.21 ). Near the luminal surfaces of the cells, smaller lateral processes extend from the primary ridges to interdigitate with those of adjacent cells. Small basal villi that do not contain mitochondria are found along the basal cell surfaces ( Figs. 2.18, 2.19, 2.21, and 2.22 ). These extensive interdigitations result in a complex extracellular compartment, the basolateral intercellular space ( Figs. 2.21–2.23 ), which is separated from the tubule lumen (apical cell surface) by the tight junctions (zonula occludens). Proximal tubule tight junctions express specific claudin proteins, which confer specific ion permeabilities and likely contribute to the high rates of paracellular sodium and water transport. Below the tight junction lies the beltlike intermediate junction, the zonula adherens, followed by several desmosomes distributed randomly at variable distances beneath the intermediate junction. In mammalian and invertebrate renal proximal tubules, gap junctions are present in small numbers and can provide a pathway for the movement of ions between cells and for cell–cell communication via a family of proteins known as connexins. The lateral intercellular space of each PCT cell is open at the basement membrane, which separates the cell from the peritubular interstitium and capillaries. The thickness of the basement membrane gradually decreases along the proximal tubule. For example, in the rhesus monkey, the basement membrane thickness is approximately 250 nm, 145 nm, and 70 nm in the S1, S2, and S3 segments, respectively.

Schematic drawing illustrating the three-dimensional configuration of the proximal convoluted tubule cell.

From Welling LW, Welling DJ. Shape of epithelial cells and intercellular channels in the rabbit proximal nephron. Kidney Int. 1976;9:385–394.

Scanning electron micrograph of rat proximal convoluted tubule, illustrating the lush brush border (BB) , primary cilia extending into the lumen (L) , prominent lateral cell processes, and multiple small basal processes (B) , called micropedici.

Modified from Verlander JW. Solute reabsorption. In Cunningham’s Veterinary Physiology . 6th ed. St Louis: Elsevier; 2020.

Transmission electron micrograph of the proximal convoluted tubule from a normal human kidney.

The mitochondria (M) are elongated and tortuous, occasionally doubling back on themselves. The endocytic apparatus, composed of apical vacuoles (AV), apical vesicles (V), and apical dense tubules (arrows), is well developed. G, Golgi apparatus; IS, intercellular space; L, lysosome; Mv, microvilli forming the brush border; TL, tubule lumen.

The lateral cell processes of PCT cells combined with extensive invaginations of the plasma membrane increase both the intercellular space and surface area of the basolateral plasma membrane. In rabbits, the area of the lateral surface equals that of the luminal surface and amounts to 2.9 mm ² per mm of tubule. Elongated mitochondria are located in the lateral cell processes near the plasma membrane (see Figs. 2.18 and 2.23 ), where sodium–potassium adenosine triphosphatase (Na ⁺ -K ⁺ -ATPase) resides. ^, Although mitochondria often appear rod-shaped in two-dimensional images, many mitochondria are branched and connected with one another. A system of smooth membranes, the paramembranous cisternal system, which may be in continuity with the smooth endoplasmic reticulum, is often observed between the plasma membrane and mitochondria. PCT cells contain large quantities of smooth and rough endoplasmic reticulum, and free ribosomes are abundant. A well-developed Golgi apparatus, composed of smooth-surfaced sacs or cisternae, coated vesicles, uncoated vesicles, and larger vacuoles, is located above and lateral to the nucleus. In addition, an extensive system of microtubules is located throughout the cytoplasm of proximal tubule cells.

PCT cells have lush luminal brush borders formed by densely packed, fingerlike projections of the apical plasma membranes, the microvilli. The brush border greatly increases the apical cell surface, increasing the absorptive surface facing the luminal fluid. Each microvillus contains 6 to 10 actin filaments of approximately 6 nm in diameter that extend variable distances into the cell body. A network of filaments containing myosin and spectrin, the terminal web, is located in the apical cytoplasm just beneath and perpendicular to the microvilli. Each PCT cell has a well-developed endocytic–lysosomal apparatus that is involved in the reabsorption of macromolecules from the ultrafiltrate and their degradation. ^, The endocytic compartment includes an extensive system of coated pits, small coated vesicles, apical dense tubules, and larger endocytic vacuoles without a cytoplasmic coat ( Fig. 2.24 ). The coated pits are invaginations of the apical plasma membrane at the base of the microvilli and contain clathrin, megalin, and cubilin, proteins that are involved in receptor-mediated endocytosis. The cytoplasmic coat of the small vesicles is similar in ultrastructure to the coat that is present on the cytoplasmic side of the coated pits.

Transmission electron micrograph of the apical region of a human proximal tubule, illustrating the endocytic apparatus including coated pits (Cp), coated vesicles (Cv), apical dense tubules (Dat), and endosomes (E).

PCT cells contain numerous lysosomes of variable size, shape, and ultrastructural appearance ( Fig. 2.25 ). ^, Lysosomes are membrane-bound, heterogeneous organelles that contain proteases, lipases, glycosidases, and acid hydrolases, including acid phosphatases. Lysosomes degrade material absorbed by endocytosis (heterophagocytosis) and often contain electron-dense deposits that are believed to represent reabsorbed substances such as proteins (see Figs. 2.19 and 2.25 ). Lysosomes also participate in the normal turnover of intracellular constituents by autophagocytosis, and autophagic vacuoles containing fragments of cell organelles are often seen in PCT cells. Lysosomes containing nondigestible substances are called residual bodies; these can empty their contents into the tubule lumen by exocytosis. Multivesicular bodies (MVBs), which are part of the vacuolar-lysosomal system, are often observed in the cytoplasm of PCT cells. MVBs were originally thought to be involved in membrane retrieval and/or membrane disposal, but later studies suggest that MVBs may provide an exit route for plasma membrane vesicles formed by endocytosis and could function as a signaling mechanism to downstream nephron segments. ^, The extensive vacuolar-lysosomal system of proximal tubule cells plays an important role in the reabsorption and degradation of albumin and low-molecular-weight plasma proteins from the glomerular filtrate. ^{, ,} Under normal conditions, the vacuolar-lysosomal system is most prominent in the PCT, but in proteinuric states, large vacuoles and extensive lysosomes can be observed in the PST as well. ^,

Transmission electron micrographs illustrating the appearance of different types of lysosomes from human proximal tubules.

(A) Lysosomes. Several mitochondria (M) are also shown. (B) Early stage of formation of an autophagic vacuole. (C) Fully formed autolysosome containing a mitochondrion undergoing digestion. (D) Autolysosome containing a microbody undergoing digestion. A multivesicular body (arrow) is also shown.

From Tisher CC, Bulger RE, Trump BF. Human renal ultrastructure. I: proximal tubule of healthy individuals. Lab Invest. 1966;15:1357–1394.

In the rat, the proximal straight tubule (PST, pars recta) includes the terminal portion of the S2 segment, located in the medullary ray, and the entire S3 segment. PST morphology varies considerably among species. For example, the rat S3 brush border measures up to 4 μm long, whereas in the rabbit and human PST, the brush border is relatively short. The S3 epithelium is simpler than both the S1 and S2 segments. ^, Basolateral plasma membrane invaginations are virtually absent, mitochondria are small and randomly scattered throughout the cytoplasm, and intercellular spaces are smaller and less complex ( Figs. 2.20 and 2.26 ). These morphologic characteristics are in agreement with studies demonstrating that Na ⁺ -K ⁺ -ATPase activity is significantly less in the PST compared with PCT. In contrast to PCT cells, the vacuolar-lysosomal system is less prominent in rat S3 cells, although in both rabbits and humans, many small lysosomes containing electron-dense membranelike material are present in the late PST. ^{, ,} Peroxisomes are common in the PST ( Fig. 2.27 ). In contrast to lysosomes, peroxisomes are irregular in shape, are surrounded by a 6.5-nm-thick membrane, and do not contain acid hydrolases. Peroxisomes within the PST vary considerably in appearance among species. In the rat, small, circular profiles are visible by transmission electron microscopy just inside the limiting membrane, and rod-shaped structures often project outward from the organelle. In addition, a small nucleoid is often present in peroxisomes in the PST. Peroxisomes contain abundant catalase, which is involved in the degradation of hydrogen peroxide, and various oxidative enzymes including l-α-hydroxy-acid oxidase and d -amino acid oxidase. ^,

Low-magnification transmission electron micrograph of a segment of the proximal straight tubule from a human kidney.

The microvilli on the convex apical cell surface are not as long as those in the rat proximal straight tubule. The lysosomes are extremely electron dense. The clear, single membrane–limited structures at the base of the cell to the right represent lipid droplets.

Courtesy R.E. Bulger, Ph.D.

Transmission electron micrograph of the rat proximal straight tubule, S3 segment.

Endocytic vesicles, lysosomes, and autophagic vacuoles (arrowhead) are less abundant than in S1 and S2 segments. However, peroxisomes are abundant and identified by their irregular, angular shape and small, circular protuberances along the edges (arrows) .

The proximal tubule plays a major role in the reabsorption of Na ⁺ , HCO ₃ ⁻ , Cl ⁻ , K ⁺ , Ca ²⁺ , PO ₄ ³⁻ , water, and organic solutes such as vitamins, glucose, and amino acids; secretion of protons, ammonia, and organic anions; and uptake of filtered peptides and proteins. The ultrastructural features of proximal tubule cells aid in these transport processes, most notably the high surface density of both the apical and basolateral plasma membrane compartments, high mitochondrial density in early proximal tubule segments, and abundant endocytic vesicles and lysosomal system. Proximal tubule cells alter transport capacity in some instances by redistribution of specific transporters located in the brush border. For example, the apical sodium-hydrogen exchanger, NHE3, is rapidly redistributed between the brush border microvilli and base of the microvilli in models that alter proximal tubule sodium uptake, ^, whereas parathyroid hormone stimulates redistribution of the sodium phosphate transporter, NaPi-2, from the microvilli to endosomes. Changes in the hydraulic and oncotic pressures across the tubule and capillary wall cause significant ultrastructural changes in the proximal tubule, especially in the configuration of the lateral intercellular spaces. ^,

The TAL arises abruptly from the thin limbs of loop of Henle, spans the inner and outer stripes of the outer medulla, extends through the cortex in the medullary rays, contacts the glomerulus of its own nephron at the macula densa, and extends a short distance beyond the macula densa before the transition to the DCT ^, (see Fig. 2.5 ). In short-looped nephrons, the transition to the TAL can occur shortly before the hairpin turn, but this is not the case in all species. In the outer medulla, TAL cells are taller in the inner stripe, beginning at ∼11 μm and declining to between 7 and 8 μm in height. ^{, ,} As the tubule ascends toward the cortex, cell height gradually decreases further to ∼5 μm in the cortical TAL (cTAL) of the rat. In rabbits, the cTAL is lower than the medullary TAL (mTAL) in height, averaging 4.5 μm but declining to ∼2 μm in the terminal part. ^,

TAL cells have extensive infoldings of the basolateral plasma membrane and interdigitations between adjacent cells ( Figs. 2.33 and 2.34 ). The basolateral infoldings often extend from the base to two-thirds or more of the cell height, particularly in the mTAL in the inner stripe. The cell nucleus is centrally located, with little cytoplasm or organelles between the nucleus and either the apical or basal surface. Abundant elongated mitochondria are located in lateral cell processes, generally oriented perpendicularly to the basement membrane, similar to the S1 segment of the proximal tubule, and they contain prominent granules in the matrix. The TAL also has a well-developed Golgi complex; small, subapical cytoplasmic vesicles and tubulovesicles; multivesicular bodies and lysosomes; and abundant smooth and rough endoplasmic reticulum. The tight junctions are 0.1 to 0.2 μm in depth in the rat ; in rabbit, the length of the tight junctions increases from the mTAL to the cTAL. Intermediate junctions are also present, but desmosomes appear to be lacking.

Scanning electron micrograph illustrating the luminal surface of rat medullary thick ascending limb.

The white asterisk denotes smooth-surfaced cells; the black asterisk identifies rough-surfaced cells.

Modified from Madsen KM, Verlander, Tisher CC. Relationship between structure and function in distal tubule and collecting duct. J Electron Micros Tech. 1988;9:187–208.

Transmission electron micrograph of cortical thick ascending limb (TAL) in rat kidney.

The apical surface has numerous short apical microprojections, typical of the rough TAL cells common in the cortical TAL, and deep, complex invaginations of the basal plasma membrane extend into the apical region of the cell and enclose elongated mitochondrial profiles. There are few organelles and little cytoplasm between the nucleus and the apical and basal aspects of the cell compared with the distal convoluted tubule (see Fig. 2.36 ). A peritubular capillary with fenestrated endothelium (arrows) is adjacent to the basal side of the TAL cell.

By SEM, the TAL of the rat kidney has two morphologically distinct cells, designated “smooth” and “rough,” distinguished by the appearance of the luminal plasma membrane and lateral cell borders. Rough TAL cells have numerous small apical microprojections, whereas the apical surface of smooth TAL cells has few microprojections except along the cell borders (see Fig. 2.33 ); both types have a single, central primary cilium. In the inner stripe of the outer medulla, rough TAL cells generally have prominent lateral processes that interdigitate with neighboring cells, producing an undulating cell border, whereas smooth TAL cells typically have only shallow lateral processes and hence a relatively simple cell border; these differences are not present in the cTAL. However, compared with rough TAL cells, smooth TAL cells have a more prominent subapical cytoplasmic vesicle and tubulovesicle compartment. The smooth surface pattern predominates in the mTAL, but as the thick limb ascends, the number of rough TAL cells increases, and luminal microprojections and apical lateral invaginations become more prominent. Consequently, the surface area of the luminal plasma membrane is significantly greater in the cTAL than in the mTAL.

The structural characteristics of the TAL, notably the high density of mitochondria interposed between extensive basolateral plasma membrane infoldings, contribute to its important role in active reabsorption of NaCl via the Na ⁺ , K ⁺ , 2Cl ^– cotransporter located in the apical plasma membrane driven by abundant basolateral Na ⁺ -K ⁺ -ATPase. Within the TAL, axial heterogeneity in the ultrastructural features correlates with Na ⁺ -K ⁺ -ATPase activity, with the mTAL in the inner stripe having the greatest basolateral plasma membrane area, mitochondrial density, and Na ⁺ -K ⁺ -ATPase activity. ^{, , ,} However, functional correlations with the observed axial and cellular structural heterogeneity are limited. Some physiologic studies using the isolated perfused tubule technique found that NaCl transport is greater in the medullary segment than in the cortical segment of the TAL, consistent with the structural differences, but others did not observe this. ^, Similarly, functional differences between the smooth and rough forms of TAL cells have not been defined. Although cellular heterogeneity in the expression of various transport proteins, including ROMK, H ⁺ ATPase, and NKCC2, has been observed in the TAL, these variations in protein expression have not yet been correlated with the apical surface patterns described using electron microscopy.

The abrupt transition from the TAL to the DCT occurs a short distance distal to the macula densa and is located in the cortical labyrinth ( Figs. 2.5 and 2.35 ). Like TAL cells, DCT cells contain extensive basolateral plasma membrane infoldings and a dense array of long mitochondria aligned with the plasma membrane infoldings perpendicular to the basement membrane ( Fig. 2.36 ). However, DCT cells are significantly taller than TAL cells, and the cell nuclei are close to the apical plasma membrane with basolateral plasma membrane infoldings and mitochondria interposed between the nucleus and basement membrane. On the luminal surface, DCT cells have a single central cilium and numerous small microprojections, which are more prominent at the lateral cell borders; the lateral borders are simple compared with TAL cells ( Fig. 2.37 ). The junctional complex is composed of a tight junction, which is approximately 0.3 μm in depth, and an intermediate junction. The Golgi complex is well developed, and lysosomes and multivesicular bodies are present but less common than in the proximal tubule. The cells contain numerous small subapical vesicles, microtubules, free ribosomes, and rough and smooth endoplasmic reticulum.

Micrographs depicting the abrupt transition (arrows) from the thick ascending limb of Henle (below) to the distal convoluted tubule (above) .

(A) Light micrograph of normal rat kidney. (B) Scanning electron micrograph of normal rabbit kidney.

B, Courtesy Ann LeFurgey, Ph.D.

Transmission electron micrograph of rat distal convoluted tubule (DCT).

Although the structure of DCT cells is similar to thick ascending limb (TAL) cells in many ways, DCT cells are considerably taller, with numerous basal plasma membrane infoldings and mitochondria interposed between the nucleus and the basement membrane. Compare with the TAL cell in Fig. 2.34 .

Scanning electron micrograph showing the luminal surface of a distal convoluted tubule from a rat kidney.

Short microvilli are prominent, cell borders are accentuated by longer and more abundant microvilli, and the cell borders in the apical region are simple, lacking the interdigitations seen in the thick ascending limb. (Compare with Fig. 2.33 .)

As mentioned previously, in micropuncture studies, the “distal tubule” includes tubule segments from immediately distal to the macula densa to the first junction with another renal tubule, which includes as many as four different types of epithelia ( Fig. 2.38 ). In general, the “early” or “bright” distal tubule corresponds largely to the DCT plus a short segment of TAL, whereas the “late” or “granular” distal tubule corresponds to the connecting tubule and the initial portion of the collecting duct in the cortical labyrinth, the ICT. ^, In several species, the DCT exhibits axial heterogeneity with respect to cell morphology and transporter expression. In rabbits, the transition from DCT to CNT is morphologically distinct, but in rats, mice, and humans, the late portion of the DCT shares features of the CNT, including the presence of intercalated cells and several proteins expressed in CNT cells. In fact, two DCT segments, DCT1 and DCT2, have been defined in the rat on the basis of protein expression characteristics. Expression of the apical thiazide-sensitive NaCl cotransporter, NCC, is definitive for DCT cells and is present throughout the entire DCT. The initial DCT segment, DCT1, is distinguished from the late segment, DCT2, by the presence in DCT2 of the Na ⁺ –Ca ²⁺ exchanger (NCX1) and vitamin D–dependent calcium-binding protein, calbindin-D28K, proteins that are also expressed in CNT cells. In mice ^, and humans, NCX and calbindin-D28K are expressed throughout most of the DCT; thus DCT1 and DCT2 are not distinguishable, at least not as originally defined. Nonetheless, in rats, mice, and humans, the early DCT is distinct from the late DCT in that the latter portion expresses the epithelial sodium channel (ENaC), which is also expressed in CNT cells, and in mice and rats, the late DCT expresses the apical calcium channel TRPV5, which is absent from the early DCT. ^, Furthermore, in rats and mice, the late portion of the DCT also contains intercalated cells, predominantly the so-called non-A, non-B subtype, which is described in detail in the section on the collecting duct.

Diagram of the various anatomic arrangements of the distal tubule and cortical collecting duct in superficial and juxtamedullary nephrons.

(See text for detailed explanation.) CCD, Cortical collecting duct; CNT, connecting segment; DCT, distal convoluted tubule; G, glomerulus; ICT, initial collecting tubule; MD, macula densa; TAL, ascending thick limb (of Henle).

The DCT has the highest Na ⁺ -K ⁺ -ATPase activity of all nephron segments, ^, which drives ion transport and correlates with the high mitochondrial density and elaborate basolateral plasma membrane infoldings in this segment. The NCC is present in the apical plasma membrane and subapical vesicles. A number of studies have demonstrated structural changes in the DCT in response to physiologic stimuli that alter the transport activity in this segment. For example, treatment with furosemide, an inhibitor of the TAL transporter NKCC2, causes a marked increase in DCT cell size, basolateral plasma membrane area, and cell proliferation, along with increased NCC expression and sodium uptake, suggesting structural adaptations correlating with functional adaptations to conserve sodium when NKCC2 is inhibited and NaCl delivery to the DCT is increased. ^, In animals fed a low-salt diet, NCC is largely expressed in the apical plasma membrane, where it mediates apical NaCl uptake, whereas feeding a high-salt diet or acute induction of hypertension causes redistribution of NCC to the subapical cytoplasmic vesicles. ^, Conversely, angiotensin II administration acutely causes a significant increase in the apical plasma membrane expression of NCC and a reduction in NCC expression in apical cytoplasmic membrane vesicles, whereas treatment with captopril, an angiotensin-converting enzyme inhibitor, has the opposite effect on NCC distribution, although under these conditions, phosphorylated NCC is found exclusively in the apical plasma membrane and not in cytoplasmic vesicles. Similarly, estradiol administration in ovariectomized rats, which increases NCC phosphorylation and activity, causes an increase in apical plasma membrane complexity and apical plasma membrane NCC expression along with depletion of apical cytoplasmic vesicles. Thus structural changes occur in the DCT in response to stimuli that alter NCC transporter expression and functional activity.

The CNT constitutes the main portion of the “late” or “granular” distal tubule as defined in micropuncture literature. The CNTs of superficial nephrons continue directly into ICTs, whereas CNTs from midcortical and juxtamedullary nephrons join to form arcades that ascend in the cortex and continue into ICTs (see Fig. 2.38 ). ^, CNTs are present throughout the cortical labyrinth but are present in higher density surrounding interlobular vessels; the CNT makes contact with the afferent arteriole of its own glomerulus, upstream of the JGA. ^, These contacts enable crosstalk between the CNT and renal vasculature, which regulates renal perfusion in addition to the classic tubuloglomerular feedback mechanism mediated through the JGA.

In the rabbit, CNT is a well-defined segment composed of two cell types: the CNT cell and the intercalated cell. ^, However, in other species, including rats, ^, mice, and humans, ^, the transition from DCT to CNT is not structurally distinct. More distally, rat and mouse CNT clearly differs from late DCT in the increased frequency of intercalated cells and the structural characteristics of the majority cell type, the CNT cell ( Fig. 2.39 ). The CNT transitions to the ICT, which is in the cortical labyrinth and connects the CNT to the CCD, which is located in the medullary ray. The ICT is a lower epithelium than the CNT and, like the CCD, is made up of principal cells and intercalated cells ( Fig. 2.40 ). In rabbit kidney, the CNT to ICT transition is distinct. In rat kidney, although there is some intermingling of CNT cells and principal cells in the late portion of the CNT, the CNT and ICT are largely distinguishable by morphologic characteristics of CNT cells versus principal cells. In mice, the transition from CNT to ICT is more gradual, and only the early CNT and late ICT are clearly identifiable as such based on cellular morphology.

Transmission electron micrograph of rat connecting segment (CNT) cell.

The CNT cell has deep basolateral plasma membrane infoldings with numerous mitochondria, but a lower mitochondrial density than distal convoluted tubule (DCT) cells, and the nucleus is typically rounder than DCT cell nuclei. (Compare with Fig. 2.36 .)

Light micrograph of initial collecting tubules (asterisks) in toluidine blue–stained rat kidney.

One tubule lies just beneath the renal capsule (top of picture) , where it would be easily accessible to micropuncture. Dark staining cells (arrows) are intercalated cells. This segment of the collecting duct corresponds to the so-called late distal tubule as defined in micropuncture studies.

CNT contains primarily two cell types: the majority cell type, the CNT cell, which occurs only in this segment; and intercalated cells, which account for approximately 40% of the cells. The CNT cell is tall with an apically located nucleus like the DCT cell but has a rounder nucleus, more cytoplasm and organelles between the apical plasma membrane and the nucleus, shallower and less uniform basolateral plasma membrane infoldings, and fewer, more randomly arranged mitochondria.

Three distinct intercalated cell subtypes are present in the CNT, based on not only morphologic characteristics but also distinct patterns of transporter expression: type A, type B, and non A, non B. In rat and mouse CNT, the so-called non A, non B intercalated cell is the most prevalent subtype, followed by type A cells and type B cells. The morphologic characteristics and localization of specific ion transporters distinguishing these three intercalated cell subtypes and their axial distribution are detailed in the following section.

The CNT contributes to regulated reabsorption of sodium, calcium, and water via specific proteins expressed in CNT cells. In addition to basolateral Na ⁺ K ⁺ -ATPase, CNT cells and principal cells in the ICT express the ENaC ^{, ,} and apical potassium channel, ROMK. Proteins mediating calcium transport expressed by CNT cells include the basolateral Na ⁺ -Ca ²⁺ exchanger, NCX1, Ca ²⁺ -ATPase, apical TRPV5, and calbindin-D28K. ^, In rats, mice, and humans, the CNT, like the collecting duct, expresses the vasopressin-sensitive AQP2, ^{, ,} although AQP2 appears to be absent from the rabbit CNT.

In rat kidney, a small population of cells in the late DCT at the transition to the CNT expresses both the NCC and the Na ⁺ -Ca ²⁺ exchanger, which have been considered specific for DCT cells and CNT cells, respectively. ^, In rabbit kidney, the CNT is distinct from the DCT in both structure and function, and cells coexpressing NCC and the Na ⁺ -Ca ²⁺ exchanger are not evident in either segment. The CNT makes a transition to the ICT, a segment composed of principal cells and intercalated cells located in the cortical labyrinth before joining with the CCD in the medullary ray. Expression of the Na ⁺ -Ca ²⁺ exchanger is abundant in the basolateral plasma membrane of CNT cells but undetectable in CCD principal cells.

Potassium loading in rats stimulates potassium secretion in the CNT and increases the basolateral plasma membrane surface area of CNT cells and principal cells of both the CNT and the ICT, where Na ⁺ -K ⁺ -ATPase resides, as well as the apical expression of the potassium channel, ROMK. In rabbits, feeding a low-sodium, high-potassium diet induces a similar increase in CNT basolateral plasma membrane. These structural changes are consistent with increased basolateral Na ⁺ -K ⁺ -ATPase expression and activity driving apical potassium secretion.

The collecting duct in the cortex includes ICT in the cortical labyrinth and the CCD in the medullary ray (see Fig. 2.5 ). The cells of the ICT are taller than those of the CCD and generally have more complex plasma membrane microprojections and infoldings ( Fig. 2.42 ) and more intense plasma membrane transporter expression, but otherwise these two subsegments are morphologically similar. Nonetheless, because of these subtle differences in ultrastructure, it is important to distinguish between ICT and CCD when quantifying morphologic components or subcellular immunolabeling.

Transmission electron micrograph of a principal cell from the initial collecting tubule (ICT) from normal rat kidney.

Principal cells in the ICT are similar to those in the cortical collecting duct but typically slightly taller with more extensive infoldings of the basal plasma membrane.

Principal cells account for approximately 60%–65% of cells in the CCD of rabbit, as in rat and mouse. ^{, ,} The nucleus in principal cells is located close to the apical surface, a feature that helps distinguish principal cells from intercalated cells, which have a central or basal nucleus. By transmission electron microscopy, principal cells have relatively few small cytoplasmic vesicles between the nucleus and the apical plasma membrane, a light-staining cytoplasm, and few apical plasma membrane microprojections. By SEM, these apical microprojections appear as short, stubby microvilli that are less numerous near the single central cilium than at the periphery of the cell (see Fig. 2.41 ). Principal cells have numerous basal plasma membrane infoldings, without interposition of mitochondria or other organelles (see Fig. 2.42 ). Lateral cell processes and interdigitations are virtually absent. Cell organelles are relatively sparse: Mitochondria are small and scattered randomly in the cytoplasm; there are few lysosomes, autophagic vacuoles, and multivesicular bodies; and the Golgi body, rough and smooth endoplasmic reticulum, and free ribosomes are present but not prominent.

Intercalated cells in the CCD are distinguishable from principal cells by several morphologic features. As mentioned earlier, intercalated cells lack cilia and have a more centrally or basally located nucleus versus the apical nucleus of principal cells. Under basal conditions, intercalated cells have more abundant apical plasma membrane microprojections, higher mitochondrial density, abundant cytoplasmic vesicles, numerous polyribosomes, and a prominent Golgi apparatus. In the CNT through CCD, intercalated cells in plastic sections stain more intensely with toluidine blue than principal cells, partly due to cytoplasmic staining and partly due to the high mitochondrial density. This characteristic earned them the designation “dark cells,” in early morphologic studies (see Fig. 2.40 ). By transmission electron microscopy, the electron density of intercalated cell cytoplasm varies with the subtype but is generally somewhat darker than surrounding principal and CNT cells.

In the rat, mouse, and human kidney, three distinct intercalated cell subtypes are present in the CNT, ICT, and CCD. These subtypes are recognizable by ultrastructural features and cell-specific expression and subcellular distribution of various membrane and cytoplasmic proteins. Ultrastructural studies characterized two distinct populations of intercalated cells in the rat CCD, types A and type B, with ∼60% of intercalated cells identified as type A and ∼40% as type B ( Fig. 2.43 ). The existence of a third distinct intercalated cell subtype was recognized later, the so-called non-A, non-B intercalated cell, which occurs almost exclusively in the CNT and ICT ( Fig. 2.44 ). ^{, ,}

Transmission electron micrograph from rat cortical collecting duct illustrating type A (right) and type B (left) intercalated cells under basal conditions.

Note differences in the density of the cytoplasm, location of the nuclei, distribution of the mitochondria and cytoplasmic vesicles, and number of apical projections between the two cell types.

Modified from Madsen KM, Verlander JW, Tisher CC. Relationship between structure and function in distal tubule and collecting duct. J Electron Microsc Tech. 1988;9:187–208.

Transmission electron micrograph of a non-A, non-B intercalated cell in rat kidney.

This intercalated cell subtype has a high mitochondrial density compared with type A and type B intercalated cells, complex basolateral plasma membrane infoldings, and abundant long apical microprojections under basal conditions.

By transmission electron microscopy under basal conditions, type A intercalated cells are characterized by moderate apical plasma membrane microprojections, a prominent apical cytoplasmic vesicle compartment, numerous mitochondria, a centrally located nucleus, and moderate basolateral plasma membrane infoldings (see Fig. 2.43 ). Profiles of the subapical vesicles appear as spherical vesicles and elongated tubulovesicles, which occasionally can be seen in contact with, or invaginating from, the apical plasma membrane ( Fig. 2.45 ). Both the apical plasma membrane and the apical cytoplasmic vesicles are relatively electron-dense, in part due to coating of the cytoplasmic surfaces with characteristic club-shaped particles or “studs,” which are also present in the structurally similar OMCD intercalated cells ^, and which are associated with the vacuolar proton pump, H ⁺ -ATPase. By SEM, the apical plasma membrane microprojections are mostly in the form of small folds, “microplicae,” rather than microvilli, again similar to OMCD intercalated cells. ^{, ,}

Transmission electron micrograph illustrating the apical region of a type A intercalated cell from a rat kidney.

Note the large number of tubulovesicles (solid arrows) , invaginated vesicles (open arrows) , and small coated vesicles with the appearance of clathrin vesicles (arrowheads) .

By SEM under basal conditions, type B intercalated cells typically have a smaller, more angular luminal cell outline than type A cells and only sparse microprojections, mostly in the form of short microvilli (see Fig. 2.41 ). By transmission electron microscopy, the type B intercalated cell has a denser cytoplasm and more abundant, frequently clustered mitochondria, and the cell nucleus is typically eccentric, rather than centered (see Fig. 2.43 ). Numerous vesicles are present throughout the cytoplasm, but the cytoplasmic vesicle membranes are more delicate in appearance and less electron dense compared with those in type A cells, and few membrane “studs” are evident. The apical plasma membrane is relatively smooth, with small, short microprojections, and a band of dense cytoplasm without organelles or vesicles typically is present just beneath the apical plasma membrane. The basolateral plasma membrane infoldings are more elaborate than in type A cells, except in regions of cytoplasmic extensions filled with vesicles that frequently contact the basement membrane. In the rat under basal conditions, the surface density of the basolateral plasma membrane in type B intercalated cells is significantly greater, and that of the apical plasma membrane is significantly less than in type A cells.

The so-called non-A, non-B intercalated cell represents approximately half of the intercalated cells in the mouse CNT and ICT but is less common in the rat. This cell was initially dubbed “non-A, non-B” because it exhibited protein expression patterns different from the recognized A and B cell types, but it was unclear whether it was a distinct cell type. It now appears clear that this cell is indeed a distinct cell type that can be characterized and differentiated from A and B cells by specific protein expression patterns, in addition to its structural features ( Figs. 2.44 and 2.46 ).

Characteristic immunolabeling of three distinct intercalated cell subtypes in the connecting segment (CNT, top panels ) and cortical collecting duct (CCD, bottom panels ) by differential interference contrast microscopy (DIC).

Type A intercalated cells express the basolateral anion exchanger, AE1; apical H ⁺ -ATPase; and basolateral ammonia transporter, Rhbg. Type B intercalated cells express the apical anion exchanger, pendrin, and basolateral H ⁺ -ATPase but no AE1 or Rhbg. The third type, the so-called non-A, non-B intercalated cell (or type C intercalated cell), expresses apical pendrin and basolateral Rhbg but no AE1. Left column: Double labeling for AE1 (brown), which is definitive for type A intercalated cells, and apical pendrin (blue), which is present in type B and non-A, non-B intercalated cells. Pendrin labeling is exclusively in AE1-negative cells. Middle column: Double labeling for AE1 (brown) and the a4 subunit of H ⁺ -ATPase (blue) . Type A intercalated cells (AE1-positive) have apical H ⁺ -ATPase label. Type B intercalated cells have basolateral H ⁺ -ATPase label (arrows) , as well as diffuse apical label, which correlates with cytoplasmic vesicle labeling shown by immunogold electron microscopy. Type B intercalated cells are uncommon in the connecting segment (CNT) but represent virtually all non-A intercalated cells in the cortical collecting duct (CCD). In the CNT, most non-A intercalated cells have apical H ⁺ -ATPase label but no basolateral label. These are non-A, non-B (type C) intercalated cells. Right column: Double labeling for pendrin (blue) and Rhbg (brown) . Type B intercalated cells and non-A, non-B intercalated cells, both pendrin-positive, can be discriminated by basolateral Rhbg expression. Non-A, non-B intercalated cells, which express basolateral Rhbg (arrowheads), are the predominant pendrin-positive cell type in the CNT. Type B intercalated cells do not express detectable Rhbg (arrows) and comprise virtually all pendrin-positive cells in the CCD. Rhbg immunolabel is also present in type A intercalated cells (open arrows), CNT cells, and CCD principal cells.

In rabbit kidney, intercalated cells are generally like those of rat and mouse. Early studies described “light” and “dark” forms of intercalated cells, with the dark form predominantly in the cortex and the light form predominantly in the outer medulla, suggesting that “light” and “dark” forms may correspond to type A and type B cells, respectively. Four different surface configurations by SEM were described in rabbit collecting duct on the basis of the presence of microplicae, long and short microvilli, and combinations of these. The precise correlation of the surface configurations to type A and B intercalated cells is not known. However, cells with microplicae were most prevalent in the outer medulla and inner portion of the CCD ; this distribution and similarity to the morphology of type A intercalated cells of rat suggest they are type A cells in rabbits as well.

It is well established that intercalated cells contribute to acid-base homeostasis by transport of protons, bicarbonate, and ammonia. All intercalated cell subtypes express carbonic anhydrase type II (CA II) throughout the cytoplasm, although the different subtypes exhibit varied patterns of expression. ^{, ,} Type A intercalated cells have the most intense immunolabeling for CA II, with more intense labeling near the apical and basolateral plasma membrane domains. CA II expression in type B intercalated cells is diffuse and relatively weak, whereas the level in non-A, non-B cells is intermediate.

The electrogenic proton pump, H ⁺ -ATPase, is also strongly expressed in all intercalated cells, but the subcellular distribution determined by immunogold electron microscopy varies among the subtypes. ^, In type A intercalated cells, H ⁺ -ATPase is present in the apical plasma membrane and apical cytoplasmic vesicle membranes. In type B intercalated cells, H ⁺ -ATPase is present in the basolateral plasma membrane but not in the apical plasma membrane, and in cytoplasmic vesicles throughout the cell, including subapical vesicles, which likely account for cells with “bipolar” H ⁺ -ATPase immunolabeling, have been observed by light microscopy ( Fig. 2.46 ). Like A cells, non-A, non-B intercalated cells have H ⁺ -ATPase in the apical plasma membrane and subapical cytoplasmic vesicles. These distinctions in H ⁺ -ATPase distribution may not be discernible by light microscopy techniques, depending on the resolution of the method and the characteristics of the anti-H ⁺ -ATPase antibody used (see Fig. 2.46 ). In rabbits, cells with the subcellular distribution of H ⁺ -ATPase typical of type A and type B intercalated cells are present in the CNT, ICT, and CCD, but in CCD under basal conditions, the most prevalent distribution pattern is exclusively in intracytoplasmic vesicles in small intercalated cells with relatively uncomplicated plasma membrane compartments, suggestive of an inactive cell. Intercalated cells in the rabbit CNT and ICT have the typical polarized distribution of H ⁺ -ATPase seen in rat and mouse intercalated cells.

Expression of the Cl ^– /HCO ₃ ^– anion exchanger, AE1, is definitive for type A intercalated cells because it is the only renal epithelial cell that expresses this protein. In rat and mouse kidney, AE1 is almost entirely expressed in the basolateral plasma membrane, ^{, , ,} whereas in rabbit kidney under basal conditions, a large portion of AE1 is intracellular in multivesicular bodies and cytoplasmic vesicles, in addition to the basolateral plasma membrane. ^,

In addition to being devoid of AE1, type B and non-A, non-B intercalated cells are distinguished from type A cells by the expression of the apical Cl ^– /HCO ₃ ^– exchanger, pendrin (Slc26a4). ^{, ,} In both type B and non-A, non-B intercalated cells, pendrin is present in the apical plasma membrane and subapical cytoplasmic vesicles, although in basal conditions, the subcellular distribution is significantly different; pendrin is predominantly in the apical plasma membrane of non-A, non-B cells but predominantly in subapical vesicles in type B cells, with little apical plasma membrane expression.

The three recognized intercalated cell subtypes also have cell-specific expression of other transporters and enzymes, particularly those involved in ammonia metabolism. Type A and non-A, non-B intercalated cells express the ammonia transporters, Rhbg and Rhcg, and cytoplasmic glutamine synthetase. ^, In both cell types, Rhbg is exclusively in the basolateral plasma membrane, whereas Rhcg is expressed in both the apical and basolateral plasma membranes in type A cells, in the apical plasma membrane in non-A, non-B cells, and in apical cytoplasmic vesicles in both cell types. ^{, ,} In contrast, type B intercalated cells do not express detectable Rhcg, Rhbg, or glutamine synthetase. ^{, ,}

The pattern of transporter and enzyme expression in intercalated cell subtypes combined with physiologic studies and morphologic studies documenting cell-specific alterations in ultrastructural features and transporter distribution in response to physiologic maneuvers together have established that type A intercalated cells secrete acid, whereas type B intercalated cells secrete bicarbonate, indicating that the existence of these functionally distinct intercalated cells subtypes is responsible for the ability of the CCD to accomplish both net acid secretion and net bicarbonate secretion. ^, Many structural studies have documented intercalated cell subtype-specific changes in ultrastructure and subcellular distribution of ion and ammonia transporters in response to physiologic disturbances. ^{, , ,} Such studies have demonstrated an increase in apical plasma membrane surface area and diminished apical cytoplasmic vesicles and redistribution of H ⁺ -ATPase and Rhcg to the apical plasma membrane in type A intercalated cells in models of acidosis ^{, ,} and increased basolateral plasma membrane surface area along with redistribution of AE1 from intracellular compartments to the basolateral plasma membrane in type A cells of acid-loaded rabbits, consistent with activation of acid and ammonia secretion by type A cells during acidosis. In contrast, chloride-depletion metabolic alkalosis decreases apical plasma membrane complexity and increases the abundance of apical cytoplasmic vesicles concomitant with internalization of H ⁺ -ATPase in type A cells, suggesting inactivation of acid secretion. In type B intercalated cells, various models in mice that enhance bicarbonate secretion and chloride uptake increase apical plasma membrane surface area, decrease apical cytoplasmic vesicles, and cause redistribution of pendrin from the cytoplasmic vesicles to the apical plasma membrane. In rats, chloride-depletion metabolic alkalosis increases type B intercalated cell size, basolateral plasma membrane complexity, and basolateral plasma membrane H ⁺ -ATPase expression.

Non-A, non-B intercalated cells can secrete bicarbonate via apical pendrin and secrete protons via apical H ⁺ -ATPase, but the net effect of these processes is poorly understood because the anatomy of the CNT, where most non-A, non-B cells reside, makes in vitro studies of these specific cells extremely difficult if not impossible. Type B and non-A, non-B intercalated cells appear to have an important role in transcellular chloride reabsorption via coordination of apical pendrin-mediated Cl ^– /HCO ₃ ^– exchange and basolateral chloride exit via ClC-K2 channels. ^, In mouse models that increase pendrin activity in type B intercalated cells, described earlier, non-A, non-B cells typically exhibit increased apical plasma membrane area and pendrin expression as well, although the relative distribution of pendrin between cytoplasmic vesicles and the apical plasma membrane frequently does not change. ^,

The functions of type A, a cid-secreting, and type B, b icarbonate-secreting, intercalated cells correspond well to their names. Although changes in nomenclature can create confusion initially, because the “non-A, non-B” cell is primarily found in the C NT and is believed to have an important role in c hloride transport, designating it as a “type C” intercalated cell would be logical.

In contrast to intercalated cells, principal cells express abundant Na ⁺ K ⁺ -ATPase in the basolateral plasma membrane, the ENaC, and the potassium channel, ROMK, in the apical plasma membrane. AQP2 is present in the apical plasma membrane and cytoplasmic vesicles, as well as in the basolateral plasma membrane. ^{, ,} The ammonia transporters Rhbg and Rhcg are less abundant than in intercalated cells in the basal state but otherwise are expressed in a pattern similar to type A intercalated cells. These transporters enable principal cells to reabsorb sodium and secrete potassium, to reabsorb water when vasopressin is present, and to contribute to ammonia secretion.

Structural correlates in principal cells in cortical segments to changes in the physiologic state are typically not as dramatic as those seen in intercalated cells. In rats and rabbits, feeding a high-potassium diet ^, or treating with aldosterone or the mineralocorticoid analog deoxycorticosterone significantly increases the basolateral plasma membrane surface area of principal cells. Furthermore, models that enhance sodium reabsorption in the CCD cause redistribution and increased apical plasma membrane expression of ENaC subunits. These structural and immunolocalization studies correlate with the roles of principal cells in regulated sodium reabsorption and potassium secretion.

Like the CCD, the OMCD epithelium is made up of principal cells and intercalated cells. In rat and mouse kidney, intercalated cells represent 35%–40% of the cells in both the OMCDo and OMCDi. ^, In rabbit kidney, the percentage of intercalated cells in the OMCD is less, approximately 18%, with the percentage declining axially from the OMCDo to the deep OMCDi. However, some investigators did not observe distinct epithelial cell heterogeneity in the rabbit OMCDi based on morphologic criteria, although the cells varied in mitochondrial content, subapical vesicle abundance, and density of rod-shaped particles in the apical plasma membrane.

Principal cells of the OMCD are structurally similar to those in the CCD, although they become slightly taller, and the number of organelles and basal infoldings decreases as the collecting duct descends through the outer medulla. Like principal cells in the CCD, those in the OMCDo express apical plasma membrane ENaC and AQP2 and basolateral plasma membrane Na ^+, K ⁺ -ATPase, consistent with their function in sodium and water reabsorption. ^{, , ,} The ammonia transporters, Rhbg and Rhcg, are also expressed in a similar pattern as in CCD principal cells, with basolateral Rhbg and apical and basolateral Rhcg, consistent with a role in ammonia secretion. ^,

Most intercalated cells of the OMCD are structurally similar to type A intercalated cells of the CCD ( Fig. 2.47 ), although with more prominent apical plasma membrane microprojections and fewer apical cytoplasmic vesicles under basal conditions. They also have similar transporter expression patterns, including H ⁺ ATPase and Rhcg in the apical plasma membrane and apical cytoplasmic vesicles and AE1, Rhbg, and Rhcg in the basolateral plasma membrane. ^, In the OMCDi, the intercalated cells are taller, there are generally fewer apical cytoplasmic tubulovesicles, and the cytoplasm is less electron dense such that it is similar to that of principal cells. Type B intercalated cells, identified by pendrin expression, are generally absent, but when present are typically sparse and limited to the OMCDo near the corticomedullary junction. The presence of only the acid-secreting intercalated cell subtype in the OMCD is consistent with this segment’s ability to secrete acid and inability to secrete bicarbonate.

Transmission electron micrograph of an intercalated cell in the outer medullary collecting duct of a normal rat kidney.

The cell has a prominent tubulovesicular membrane compartment and many microprojections on the apical surface.

Modified from Madsen KM, Tisher CC. Response of intercalated cells of rat outer medullary collecting duct to chronic metabolic acidosis. Lab Invest. 1984;51:268–276.

Under basal conditions, intercalated cells in the outer medulla exhibit moderate apical plasma membrane surface microplicae and cytoplasmic tubulovesicles; the cytoplasmic face of the tubulovesicles is coated with electron-dense studlike particles ^, associated with the vacuolar H ⁺ -ATPase (see Fig. 2.45 ). After induction of acute respiratory acidosis, metabolic acidosis, or hypokalemia, the apical cytoplasmic tubulovesicles are depleted, and apical plasma membrane microplicae proliferate and protrude into the lumen, producing a marked increase in apical plasma membrane surface area ^{, , ,} and an increase in apical plasma membrane exhibiting studs on the cytoplasmic face, visible by transmission electron microscopy. ^, These ultrastructural changes coincide with increased apical plasma membrane expression of H ⁺ ATPase and Rhcg and decreased expression of these transporters in the apical cytoplasmic vesicle compartment, consistent with enhanced proton and ammonia secretion during these conditions. ^{, ,} Conversely, during chloride-depletion metabolic acidosis, apical plasma membrane surface area and H ⁺ ATPase expression decrease, whereas apical cytoplasmic vesicles and H ⁺ ATPase expression increase, consistent with reduced acid secretion in response to alkalosis. Similar responses have been documented in type A intercalated cells in the CCD. ^{, ,}

In rabbit kidney, additional structural changes are associated with acid loading involving the redistribution of expression of AE1, the basolateral Cl ^– /HCO ₃ ^– exchanger. Under basal conditions, rabbit OMCD intercalated cells contain prominent intracytoplasmic multivesicular bodies that express AE1. After acid loading, these multivesicular bodies are reduced in number and basolateral plasma membrane boundary length and AE1 expression increase, along with the development of prominent basolateral extensions that give the cells a stellate appearance by light microscopy. These findings suggest that in rabbits, acidosis stimulates not only redistribution of proton pumps from the cytoplasmic pool to the apical plasma membrane but also redistribution of intracellular AE1 to the basolateral plasma membrane, thus enabling enhanced proton secretion and bicarbonate reabsorption.