16.1 Introduction: Why “Urogenital”?

It may seem odd to consider “uro-” and “-genital” together. After all, we might see different medical providers for issues with kidneys versus reproductive anatomy. However, the “uro-” and “-genital” organ systems share a common developmental and evolutionary history, and parts of the urinary system have been co-opted into the reproductive system. Necessarily, it is difficult to talk about one without the other.

The plan for this chapter is to focus just on the excretory parts—the uro- part of urogenital. The next chapter will cover the reproductive part. Note also that when people think about waste leaving the body, they also typically think about pooping—after all, everybody poops. This is part of the digestive system, and as such it is covered in Chapter 13. When we say excretion, we really are talking about kidneys and the associated tubes.

16.2 Structure of the Excretory System

The main functions of the excretory system are waste management and dealing with water and solute balance. This is primarily done through large, paired structures called kidneys. However, each kidney is composed of up to 1 million microscopic, tubular structures called nephrons (Greek for “of the kidney”; Figure 16.1). Nephrons are the functional units of the kidney, in that all nephrons within a kidney do not need to be in working order to produce urine.

Nephrons get their fluids (filtrate) from the renal corpuscle, which is in turn composed of the glomerulus (a ball of capillaries that receive blood from the renal artery; plural: glomeruli) that is covered by a pouch-like part of the nephron called the renal capsule (sometimes known as Bowman’s capsule, glomerular capsule, or Malpighian capsule; Figures 16.1 and 16.2).

The renal corpuscle collects fluid, cations (positively charged ions), and small molecules (e.g., glucose and amino acids) that have filtered out of the capillaries of the glomerulus. This filtrate then makes its way into a tubule. The structure of the tubule depends on the vertebrate, but we will use an amniote nephron as our example here. In tetrapods, the tubule is divided into three regions—the proximal, intermediate, and distal tubule—and filtrate moves from the renal corpuscle through the tubules in that order (Figure 16.3). As filtrate passes through these three regions of the nephron, both active and passive transport are used to move molecules and water in and out of the filtrate, depending on the needs of the organism and the concentration of solutes in the surrounding kidney tissue. Each part of the nephron is responsible for moving different types of ions and molecules. Filtrate then leaves the nephron through the distal tubule and enters the collecting tubule. Collecting tubules collect filtrate from many nephrons, and final adjustments to the filtrate content are made before it is passed on for excretion. Once the filtrate reaches the end of the collecting duct and no further adjustments are made, we call it urine.

For such a tiny organ, nephrons do a lot! They are responsible for maintaining the pH of blood. They are responsible for removing toxins from the blood, such as nitrogenous waste, and releasing them through urine. Importantly, nephrons are responsible for osmoregulation, the process that maintains the water and electrolyte (solute) balance in body fluids.

16.3 Development of the Kidney

The kidneys, and therefore the nephrons, develop from mesoderm, which forms paired nephric ridges in the left and right walls of the dorsal coelom down the length of the coelom. Each ridge is divided into several segments called nephrotomes, which will eventually develop into nephrons (Figure 16.4). The medial portion of each nephrotome develops into the renal capsule, and glomeruli will develop within the capsule. The lateral portion of each nephrotome lengthens and merges with similar regions of adjacent nephrotomes to form a single nephric duct (or nephric tubule).

Understanding the embryonic development of the kidney gives us important clues to how kidneys have continued to change through vertebrate evolution. Vertebrate kidneys follow a (three-part) development sequence, where the nephron development previously described happens in three distinct phases (Figure 16.5).

The first stage is the formation of the pronephros, which happens in the anterior part of the nephric ridge. The pronephric tubules join to form a single pronephric duct. The pronephric duct develops posteriorly until it connects with the cloaca. In most vertebrates, this is a transient stage of development, and the pronephros does not persist for long. However, a small number of vertebrates retain the pronephros, including hagfishes, lampreys, lungfishes, and a few teleost species such as salmon and trout. This is sometimes called a “head kidney,” referring to its anterior position, and is additional to other kidney tissue that develops later.

The mesonephros develops next, just posterior to the pronephros. Mesonephric tubules develop as described above, but instead of merging into a brand-new common duct, they join the preexisting pronephric duct. At this point, the name of the duct is now called the mesonephric duct to reflect this developmental change.

What happens next depends on which vertebrate you’re considering. In amniotes, the caudal-most portion develops into the metanephros and the mesonephros disappears, and a gap occurs between the metanephric and mesonephric regions of the nephric ridge. As the metanephros develops, an outgrowth of the posterior mesonephric duct called the uteric bud develops and joins with the metanephric tubules. This small portion of the mesonephric duct goes through another name change to become the metanephric duct (also known as the ureter), and the metanephros becomes the adult kidney.

In vertebrates that are not amniotes, we have a different path to follow. The metanephros does not develop, nor does the uteric bud. Nephrons still develop in the posterior part of the nephric ridge that is adjacent to the mesonephric region, but there is no gap. The newly formed tubules join the mesonephric duct, and this longer mesonephric kidney is now called the opisthonephros to reflect that it is more extended than a typical mesonephric kidney. As you might have guessed by now, we also change the mesonephric duct to be called the opisthonephric duct in this case.

It is worth noting that we have painted a very broad, general picture of kidney development here. Like many areas of comparative vertebrate anatomy, this is still an active area of study. As anatomists and developmental biologists study the nephric ridge and its development in more detail and across more organisms, exceptions to the tripartite story become apparent. In many cases, there is not a neat boundary between each of the three regions discussed above. This has led to an alternative hypothesis called the holonephric concept. The Greek root holo- means “whole” or “entire”; thus holonephric indicates that the entire kidney is one organ (the holonephros, sometimes called the archinephros), and pro-, meso-, and metanephros are all regions of that holonephros that develop in sequence. The boundary between pronephros and mesonephros in particular seems to be extra fuzzy; some larvae never develop a functional pronephros (including humans, potentially), and nonfunctional pronephric cells look very similar to degrading mesonephric cells. Experiments show that the differentiation of nephric ridge cells is largely dependent on location; for example, moving cells in the cranial region of the nephric ridge to the caudal region produces cells that look just like metanephric cells. In other words, these cells lack specificity. To date, there are no genes that are specific to the development of the pronephros, for example. Rather, the same genes are active in the development of all three regions of the holonephros. However, while some vertebrates (e.g., lampreys, hagfishes, and the occasional caecilian species) develop what looks like a holonephros during their larval stages, no adult vertebrates retain a holonephros. And there are organisms, such as hagfishes, that have a clear, demarcated pronephric region.

For now, we will continue to use the tripartite concept in our discussion of the evolution of the renal system (nephrons, kidneys, and associated urinary structures). However, as with all areas of anatomy, it is possible or even likely that our future definitions and prevailing hypotheses will change as scientists continue to learn more.

16.4 Excretory System Evolution and Diversity

The story of the evolution of the excretory system is largely an environmentally driven one. Recall that one of the main purposes of this system is to manage water and solute balance. Based on what we know about the osmoregulatory strategies of nonvertebrate deuterostomes, these early vertebrates were likely osmoconformers. This means that the osmolality (the amount of solute dissolved in a solution) of the cells and fluids of these early vertebrates was likely similar to that of the surrounding salt water. The fact that nonvertebrate deuterostomes are all osmoconformers has also been used as evidence that vertebrates arose in at least a brackish environment. Among the extant vertebrates, only hagfishes, lampreys, chondrichthyans, and coelacanths still use this strategy.

However, as vertebrates expanded into freshwater and terrestrial habitats and back to the marine realm, osmoconforming was not possible. Instead, most vertebrates are osmoregulators—the osmolality of their body fluids does not match that of the environment. As such, osmoregulator excretory systems must fight against constantly losing or gaining water and electrolytes, depending on the surrounding environment.

Cyclostomes

Both hagfishes and lampreys develop a pronephros during larval development followed by a mesonephros, as we would expect from the tripartite concept we discussed. Unlike most other vertebrates, both hagfishes and lampreys retain their pronephros into adulthood. This “head kidney” is no longer functional from an excretion standpoint, although it may be active as a lymphoid organ. Instead, the opisthonephros (remember, this is an elongated mesonephros) is the functional kidney (Figure 16.6).

While the pronephros has no glomeruli, each hagfish opisthonephros (remember, this is a paired structure!) has approximately 35 large glomeruli connected to the opisthonephric duct. In fact, the glomeruli are 10 times larger than mammal glomeruli, making them particularly useful for studying renal physiology and toxicology. Once filtrate leaves the nephron, urine empties out of the opisthonephric ducts into the cloaca. As in many fishes, kidneys are not the only excretory organ that hagfishes use. Hagfishes also excrete large amounts of nitrogenous waste (ammonia, in this case) through their gills and skin. Usually, this would be a case of simple diffusion, as there should be less ammonia in the surrounding environment than in the body fluids of the hagfish, allowing ammonia to move down its concentration gradient. However, hagfishes are scavengers that eat large rotting marine animals on the seafloor. As part of the decomposition process, these carcasses are giving off large amounts of nitrogenous compounds, like urea and ammonia, that are toxic in high concentrations. Not only can hagfishes tolerate these high concentrations both internally and externally; their skin can use active transport to excrete ammonia against the concentration gradient.

Lampreys spend the majority of their lives in the larval stage, where they hang out in sediments and filter feed. The pronephros in larval lamprey have a bundle of capillaries (glomus) that act similarly to a glomerulus but serve multiple tubules that empty into a common pronephric duct. The pronephros is functional while the lamprey remains a larva, but for most lamprey species, it largely degenerates to be nonfunctional by the time they shift to the adult stage. Before the metamorphosis to an adult, the opisthonephros develops and then serves as the functional adult kidney. As we saw in the hagfish, the gills and skin are also active sites of osmoregulation and excretion.

While most lampreys are osmoconformers, 9 of the over 30 different lamprey species are anadromous—they hatch in freshwater, migrate to salt water until they’re mature enough to breed, then return back to freshwater to breed (Figure 16.7). This creates an osmoregulation problem. Lamprey body fluids, like those of any freshwater-dwelling vertebrate, have a far higher concentration of solutes than the surrounding freshwater. They are constantly taking on freshwater and losing electrolytes. Then when they get to salt water, the problem reverses—they are losing water and gaining electrolytes. That means that these anadromous lampreys have to be osmoregulators. Given the complexity of their environmental challenges, the nephrons of these anadromous lamprey are more complex than what we’ve seen in the other cyclostomes. Their collecting ducts are more complex and subdivided into proximal, intermediate, and distal sections; these sections are flexible in how they handle moving water and solutes in and out of the filtrate. The gills also need to be nimble. In freshwater, they uptake sodium and chloride ions to fight against the loss of electrolytes, but in salt water, the gills excrete sodium and chloride to keep the osmolality of the body fluids in the right place. In addition, the lamprey’s gut joins in on osmoregulation. The esophagus is capable of absorbing ions, and the intestine acts as a site of water resorption while the lampreys reside in marine environments.

Chondrichthyans

Chondrichthyan kidneys are opisthonephric, and their general shape depends on what kind of chondrichthyan you are examining, although they are generally composed of a series of irregularly shaped lobes. For example, sharks, with their more elongated bodies, have elongated kidneys that run along the length of the abdominal cavity and get wider toward the caudal end. Batoids (rays and skates) tend to have short, wide kidneys only toward the caudal end of the body. Examined in cross section, chondrichthyan kidneys are divided into two regions—a dorsolateral bundle zone, and a ventromedial sinus zone (Figure 16.8). The tubules of the nephron are even more complex than those of mammals, with at least four loops and turns and at least five different segments that meander back and forth across both the sinus and bundle zones. The sinus zone contains loops 2 and 4 of each nephron, while the bundle zone contains loops 1 and 3. The sinus zone is named after large blood sinuses that hold blood drained from the large glomeruli that sit at the boundary between the bundle and sinus zones as well as other blood vessels. The nephron loops in the sinus zone are quite separated from each other, possibly allowing room for diffusion between the filtrate and the blood in the blood sinus. The nephron loops in the bundle zone, on the other hand, are very tightly packed (thus the name “bundle zone”). Eventually, the distal tubule will grade into a collecting tubule, which will add its filtrate to a shared collecting duct. The collecting ducts pass the filtrate into the opisthonephric duct (often referred to as a ureter in sharks), which then empties urine into the cloaca.

Marine chondrichthyans are generally considered to be osmoconformers, although they tend to run just slightly hyperosmotic compared to the surrounding ocean. Unlike the hagfishes, which generally use the same ions as the ocean to maintain their internal osmolality, just under half of the osmolality of chondrichthyan fluids are due to urea and a molecule called TMAO (trimethylamine-N-oxide). Both of these molecules are readily available, as chondrichthyans produce urea instead of ammonia as a nitrogenous waste product. TMAO is also produced and counteracts urea’s tendency to destroy proteins. Interestingly, TMAO in high concentrations is toxic once digested down to trimethylamine in our digestive system. There have been cases of people eating raw Greenland shark meat and exhibiting signs of TMAO toxicity, which includes gastrointestinal distress and neurological symptoms similar to alcohol intoxication.

As we’ve seen in other fishes, kidneys are not the only osmoregulatory organ in chondrichthyans. Chondrichthyans also osmoregulate with the rectal gland (also known as the digitiform gland and the salt gland, although it is not homologous with amniotes’ salt glands). This large gland is located near the caudal portion of the digestive tract. In marine chondrichthyans, the rectal gland plays a much bigger role in salt secretion compared to the kidney and gills (in fact, the gills in marine chondrichthyans do not secrete salt). The rectal gland receives blood from the dorsal aorta via the posterior mesenteric artery. Inside the gland are thousands of tubules that are surrounded by capillary beds that receive blood from the artery. The tubules contain a series of protein channels that create the electrochemical gradient necessary to pull salt ions from the blood into the tubules, which then flow into a central lumen. The accumulated salty liquid flows from the central lumen to the rectal duct, which joins with the intestine and dumps its contents there.

When discussing the rectal gland, we purposely mentioned that the rectal gland is important in marine chondrichthyans. While most extant chondrichthyans are indeed marine, there are fully freshwater elasmobranchs, such as the rays of the Amazon River, and euryhaline species, such as bull sharks, that can tolerate a wide range of salinities from freshwater to fully marine. In both fully freshwater species, the rectal gland tends to be much smaller, as the osmoregulatory challenge is to hold on to electrolytes instead of gaining too many. However, the structure of the rectal gland is generally similar to their marine counterparts. Unlike the marine species, gills take on a much more active role in osmoregulation in freshwater and euryhaline elasmobranchs. As we would expect to see, gill tissues actively take up sodium and chloride to make up for the passive loss of these electrolytes.

Box 16.1—Bull Sharks and Migrating Salmon

Often aspects of renal physiology and anatomy are linked to where the animal lives, which largely determines whether water and ions are more likely to passively move in versus out of the animal. For example, we would expect an animal living in the desert to have anatomical and physiological adaptations to help combat water loss. However, not all animals stay within one environment for their entire lives. Some animals shift between environments depending on their life stage. We call animals that can tolerate a wide range of salinities euryhaline. A great example is the Atlantic salmon Salmo salar (Figure 16.9).

Salmon begin to mate and hatch in freshwater, and the young salmon live in freshwater for a few months before heading out to sea. As they head down rivers into estuaries, salinity gradually increases. They spend much of their adult lives in fully marine environments. As you may have gathered, this poses some challenges. Remember, freshwater fishes tend to lose salts and gain water, whereas saltwater fishes have the opposite problem. Salmon change their behavior as they move between environments—they drink water when in the ocean but do not when they’re in freshwater. Their physiology also changes. The kidneys make a lot of very dilute urine while they are in freshwater but produce far less urine that is very concentrated when the salmon is in salt water. Most incredibly, the cells in the gills (mitochondria-rich cells, also known as chloride cells) that are in charge of dealing with salts can shift the direction that they send those salt ions. In freshwater, they can use ATP to actively transport salt ions into the blood plasma; in salt water, they switch to pumping salt ions into the surrounding salt water.

Other animals may shift between aquatic environments frequently over their entire lifetime. One particularly famous example is the bull shark Carcharhinus leucas (Figure 16.10), although we should note that there are about 40 chondrichthyan species that can do this. Bull sharks are found in warm oceans across the globe and can tolerate a range of salinities from salty oceans to freshwater rivers and lakes. This means that they need to be particularly nimble as they move across these very different environments. Recall that in salt water, chondrichthyans need to retain water and export salts. However, in fresh water, they ended up gaining too much water. Bull sharks retain approximately one-third to half as much sodium, chloride, and urea in fresh water than when they are in salt water, lowering the concentration of their body fluids to slow down water gain. Urine output also increases in euryhaline sharks living in freshwater. This indicates that the gills, rectal gland, and kidneys are shifting their physiological actions to account for the change in environment.

Osteichthyans

In general, osteichthyan kidneys are opisthonephric. However, there is a lot of variation in structure and function when you compare bony fish kidneys. We will consider actinopterygian kidneys for most of this section. If we take a look at the teleosts, we see that a number of fishes—such as tilapia, salmonids, and rockfish—retain a distinct pronephros past the larval stage. These “head kidneys” generally do not participate in excretion (Figure 16.11). For example, head kidneys can have a hematopoietic function (producing blood components) or an endocrine function (see Chapter 21). Thus, it should not be a surprise that the head kidneys tend not to have nephrons. However, there are exceptions—rockfish head kidneys do contain nephrons. For those fishes that have a nonexcretory head kidney, the mesonephric portion of the opisthonephros performs the excretory duties. Thanks to their aquatic environment, most bony fishes secrete their nitrogenous waste as ammonia, as the surrounding water quickly dilutes this toxic substance.

The details of the nephron differ across species as well but are largely dependent on what environment the fish lives in, as fishes have different osmoregulatory and water-balance challenges in fresh versus salt water (Figure 16.12).

In freshwater fishes, the glomerulus is well vascularized, and the nephron is generally large because freshwater fishes typically need to excrete a lot of the water that is passively taken in due to being hyperosmotic to their environment (Figure 16.13). On the other hand, the nephrons of marine fishes tend to be smaller and less complex, and the parts that deal with inputting water into the filtrate are reduced or absent. Interestingly, there are several marine teleosts with aglomerular kidneys—they have no glomeruli! These include gulper eels, toadfish, anglerfish, and seahorses. For these species, the tubules do all the work, and the loss of glomeruli means that the tubules do not have to reabsorb as much water from the filtrate later.

Regardless of what variety of nephrons teleost fishes have, filtrate makes its way from the collecting ducts to the mesonephric duct. Teleosts also have urinary bladders that form from the mesonephric duct. Unlike human urinary bladders, teleost urinary bladders can resorb water, sodium, and chloride from urine. The intestines of teleosts are also active in osmoregulation.

Gills are an incredibly important excretory and osmoregulatory organ as well. Remember that freshwater fishes are hyperosmotic to their environment. They have to deal with the passive loss of ions and gaining of water. While freshwater fishes do pass large quantities of very dilute urine, they also have mitochondria-rich (MR) cells (also called chloride cells) in the gills that take up ions to counter those they have passively lost. Those chloride cells are also active in the gills of marine fishes, albeit in a different way. Marine fishes drink a lot of water to regain what they passively lose to the environment. However, the water they drink is salty and adds to the ions they are passively taking in through their skin. The MR cells actively excrete those extra ions.

As demonstrated here, and as you may recall from previous chapters, the bony fish story is a complex one. While actinopterygians (the ray-finned fishes) have a marine evolutionary origin, those extant actinopterygians that live in the marine realm stem from freshwater ancestors. In other words, they reinvaded the marine environment. Similarly, when we look at the history of the sarcopterygians (lobe-finned fishes), we see that while extant coelacanths stayed in salt water, the extant lungfishes are freshwater residents, but their ancestors also originated in salt water. What does all this mean? It means that we need to be careful when we are making evolutionary inferences about excretory system evolution using extant osteichthyans. Many of our living osteichthyan kidneys reflect how marine fishes made the transition to freshwater or, in some cases, how they retransitioned back to salt water.

However, there is also substantial evidence that the rhipidistians, the sarcopterygians that gave rise to the tetrapods, lived in high-salinity environments. When one considers that both terrestrial and marine osmoregulators have similar challenges—water loss to the environment—it makes sense that excretory systems from marine rhipidistians were poised to handle the similar dehydrating challenges of the terrestrial environment.

So while lungfish are not a good window into the past kidney-wise, they are interesting sarcopterygians in their own right. The kidneys of dipnoans are similar to other freshwater osteichthyans in that they have large nephrons with glomeruli and they secrete ammonia. However, the posterior nephrons are sometimes modified for sperm transport (more on that in the next chapter!). Lungfishes live in lakes and rivers that occasionally dry out, but they can handle it like champs. African lungfish will enter a dormant state called estivation, where they burrow into the mud and form a mucus cocoon around their bodies, leaving a tiny hole in the cocoon to breathe through. This stage can last for up to four years. This cocoon allows them to avoid dehydration while drastically decreasing their metabolism. How do lungfish hold on to their water, since they’re not drinking anything? Their muscle proteins produce both urea and water, they stop secreting ammonia, and the renal capsule collapses and thickens to almost shut down filtration. When they’re ready to leave estivation, the renal capsules return to normal.

Amphibians

Extant amphibian kidneys are considered to be opisthonephric, and the nephrons in all amphibians follow the general tetrapod plan described previously, which looks very similar to freshwater fishes (Figure 16.12). However, kidneys vary in morphology depending on which type of amphibian you’re examining. Amphibian kidneys are elongate, although the degree of elongation and lobation varies. For example, kidneys of salamanders and caecilians tend to be far more elongated than those in anurans. Kidneys can also differ by life stage within a species. For example, amphibians with aquatic larvae have functional pronephros and mesonephros in the larval stage. During metamorphosis, the pronephros breaks down, and the mesonephros persists as the functional kidney.

Salamander kidneys, unlike other amphibian kidneys, are usually divided into a cranial portion (the “genital kidney,” as nephrons are used for sperm transport—more on that in the next chapter; note that this is different from the kidneys of lungfishes) and a caudal portion (the “pelvic kidney”), which does the excretory work. However, testes-bearing plethodontid salamanders do not have the cranial portion at all but retain the sperm-transporting nephrons. Regardless of which salamanders you’re looking at, urine moves from the nephrons into the collecting ducts, as they do in other amphibians. The collecting ducts merge into a common collecting tube (often referred to as a ureter), which dumps urine straight into the cloaca in some families of salamanders or into a Wolffian duct that connects to the cloaca in other families.

Amphibians also possess a urinary bladder; however, it is not homologous to other urinary bladders (Figure 16.14). Amphibian urinary bladders form from a ventral outpocketing of the cloaca. Thus, urine moves from the kidneys to the cloaca to the urinary bladder, which is unique. As we saw in teleosts, the urinary bladder is used to store water and can resorb sodium and water for osmoregulation. Amphibians, like their osteichthyan ancestors, secrete ammonia as a nitrogenous waste product.

In fishes, we saw that the gills are a major osmoregulatory organ. While many adult amphibians do not have gills, amphibians that go through an aquatic larval stage do. Similar to freshwater fishes, MR cells in larval amphibian gills are primarily responsible for active transport of ions into the blood, which are lost through the skin to their freshwater environment. For the occasional salamander that does retain gills as an adult, such as mudpuppies and axolotls, it is not clear whether the gills retain their osmoregulatory function. The integument of amphibians also acts as an osmoregulatory system. As discussed in Chapter 6, the amphibian integument is quite permeable, which allows it to function in gas exchange. That same permeability also means that water and ions easily pass between the environment and amphibian bodies, although the direction of water movement depends on whether the amphibian is in a freshwater or terrestrial environment. MR cells are also active in the integument. Interestingly, the epithelium in the colon can also regulate ion balance.

Amniotes

Amniote kidneys are metanephric; the pronephros is transient and nonfunctional when it forms (it does not always form), and the mesonephros is a functional but transient embryonic kidney. The nephron is as described previously, containing a glomerulus and multiple tubule sections. In case you haven’t yet picked up on this theme, there is a lot of variation among the amniotes in terms of kidney morphology, even within specific amniote groups. Across broad groups of amniotes, this partially reflects differences in renal function, as mammals tend to make urea as their major nitrogenous waste, whereas reptiles of all sorts tend to make uric acid.

Humans (and Pigs and Rodents)

Even within a broad group of amniotes, we see variation (Figure 16.15). You are likely familiar with the bean-shaped kidneys that humans have. Several other mammals (but not other primates!) share this kidney shape with us, including rodents, pigs, and canines. The internal organization of their kidneys is similar as well, and we will use the structure of the human kidney as our example (Figure 16.16). The outermost layer of the kidney is called the renal cortex, and it surrounds the interior layer called the medulla that is responsible for urine concentration. The medulla is composed of several lobes (multilobar), which in turn are divided into renal pyramids and surrounding connective tissue called renal columns. At the narrow portion of each pyramid are bundles of collecting ducts called renal papillae. The papillae serve as a conduit to transport urine from the nephrons to the calyces (singular: calyx) of the kidney. Human kidneys have two types of calyces—minor calyces about the renal papillae, whereas the major calyces are where the minor calyces merge. The major calyces fuse into a single renal pelvis, and urine flows from there to the ureter. The ureter from each kidney joins the urinary bladder, and the urethra takes urine from the urinary bladder to exit the body.

Humans, like other mammals, have very convoluted nephrons (Figures 16.3 and 16.17). The nephron begins in the cortex, where the glomeruli are situated. As described previously, the filtrate next enters the proximal convoluted tubule, and intermediate tubule, and then the distal convoluted tubule. However, the intermediate tubule has morphologically and physiologically distinct parts, and so we give it a special name—the loop of Henle. The thin descending limb of the loop of Henle receives filtrate from the proximal convoluted tubule and extends down into the medulla; this part of the loop is highly permeable to water but not to ions or urea. After the lowest point of the loop, we move into the thin ascending limb of the loop that is only permeable to ions and moves filtrate toward the cortex. The ascending limb grades into the thick ascending limb as it continues to move toward the cortex; physiologically, its job is to remove ions from the filtrate and return them back to the plasma. Finally, the cortical thick ascending limb is where the limb enters the cortex and connects to the proximal convoluted tubule. The final components of the loop of Henle are the vasa recta and peritubular capillaries. Together, this net of arterioles, capillaries, and venules surrounds the tubule and helps maintain the concentration gradient in the kidney that allows ions and water to move across the walls of the tubules. The loops of Henle are arranged in parallel, which contributes further to the ability to make urine more concentrated than plasma.

The long loops of Henle are what allow mammals to retain water and produce very concentrated urine, which is helpful in a terrestrial environment where water loss is constant. While the loop of Henle may not dip terribly far into the medulla in many mammals, desert rodents such as kangaroo rats and spinifex hopping mice have some of the longest loops of Henle for their body size among the mammals. Their loops of Henle extend far into the medulla and have several physiological modifications, allowing them to recoup more water from the filtrate than other rodents. These desert rodents also have a whole other suite of adaptations beyond the kidney to conserve water, including being able to produce and harvest more water during cellular respiration; storing digested carbohydrates as glycogen instead of lipid, which takes less water; and producing fecal pellets that are far drier than other rodents.

Other Amniotes

Not all multilobar kidneys look like human (or pig or rodent) kidneys. Bovines have multilobar kidneys that have prominent lobes and no renal pelvis. The multilobar kidneys of cetaceans, pinnipeds, bears, hippos, and otters are even more extreme—they are reniculate, resembling a cluster of grapes (Figure 16.18). Each “grape” is called a renicule, which has its own distinct cortex, medulla, and calyx. On the other side of the spectrum, cats have unilobar kidneys, where the papillae are fused into one renal crest and there is only one pyramid per kidney. Unilobar kidneys also lack calyces, and urine passes directly into the renal crest.

We can also think about kidney shape in terms of the structure of the renal pyramids. Small mammals (smaller than rabbits) tend to have unipapillary kidneys, where the renal pyramids fuse into one papilla. Mammals that are in the size range between rabbits and kangaroos tend to have crest kidneys, which have single cortex and multiple medullary and papillary regions. However, it’s worth noting that there are some larger mammals (giraffes, camels, horses) that still have crest kidneys, and there are animals larger than rodents, such as dogs, that have unipapillary kidneys.

What has driven the diversity of mammalian kidneys, then? Unilobar kidneys are the ancestral state for mammals, and multilobar kidneys have evolved independently, multiple times. Many of the mammals with multilobar kidneys have ancestors that were either fully or semiaquatic. As we’ve already seen, the salinity of an organism’s environment (and the food in it!) affects how osmoregulation works. For mammals that are marine or have marine ancestors, having a lot of machinery (nephrons) to handle that salt intake is necessary. In addition, multilobar kidneys tend to be in larger mammals. If we consider that metabolic needs increase as mammal size increases, the number of nephrons needed also will increase. There are only so many nephrons you can stuff into one place; a multilobar kidney may have provided that extra space.

Nonavian reptiles have an even wider variety of kidney shapes than mammals, which is tied in part to body shape as we saw in the amphibians. As you may expect, snakes tend to have very long, thin kidneys, whereas lizard kidneys are shorter and more compact (Figure 16.19). Unlike mammal kidneys, and we will see next in birds, nonavian reptile kidneys are not subdivided into a medulla and cortex.

Avian kidneys are generally uniform in shape across different species, but still structurally complex (Figure 16.20). They are located in the renal fossa of the synsacrum but are still retroperitoneal like other vertebrate kidneys—they lay external to the peritoneum. Most avian kidneys can be divided into three regions—cranial, middle, and caudal—although the distinctness of each region may vary across species. Each region is composed of lobes, and each lobe has a large cortex and small medulla, as we saw in mammals.

Two different types of nephrons are found within each lobe: nephrons that more closely resemble those of nonavian reptiles, and nephrons that more closely resemble those of mammals (Figure 16.20). The nephrons that are “reptilian-type” are located in the cortex and are relatively simple with no intermediate segment, very few loops and folds, and a small glomerulus. The medullary “mammalian-type” nephrons look strikingly similar to mammalian nephrons, with a larger glomerulus and a portion that looks very similar to a loop of Henle with a thinner descending limb, thicker ascending limb, and vasa recta. And like mammalian kidneys, the mammalian-type nephrons are arranged similarly, in parallel, allowing birds to be the only other vertebrate besides mammals to make urine that is more concentrated than their plasma (although mammals can make more concentrated urine than birds).

There are also often nephrons that are somewhat intermediate between the two types of nephrons, and these are located in the transition between the cortex and medulla. All types of nephrons pass their filtrate to collecting ducts, which are located in a structure called a medullary cone that also contains the loops of Henle, and then on to the ureter. Note that there is no renal pelvis, as we saw in mammals. The ureter discharges urine into the cloaca.

Reptiles of all kinds have other means of osmoregulation alongside the renal system. The cloaca can reclaim water, sodium, and chloride. Similar to amphibians, the urinary bladders of Gila monsters are able to resorb water back into the body when food and water are not available. Birds who spend a long time at sea without access to freshwater—as well as sea turtles, sea snakes, and marine iguanas—have similar osmoregulatory challenges as marine fishes. They need to drink salt water because they’re constantly losing water to their environment, which comes with taking in extra salts. Since these amniotes do not have gills and chloride cells, they have to rely on salt glands to excrete the extra salts. These cranial glands are capable of secreting fluid that is far more concentrated than the urine’s concentration. It’s not just marine reptiles that use salt glands. Many lizards also have salt glands to keep their plasma electrolytes balanced; for example, desert iguanas use their salt glands to excrete high concentrations of potassium with minimal water loss.

16.5 Integration with Other Systems

We have already seen that several other systems work alongside the renal system to accomplish excretion and osmoregulation. In the next chapter, we will also see how tightly reproductive systems have evolved and integrated with renal systems.

We can look to humans for another function of the renal system—participating in maintaining and adjusting blood pressure. When blood pressure is too high, renal perfusion (the amount of blood flow that passes through a mass of renal tissue within a given time) rises. Simultaneously, stretch receptors in the right atrium of the heart are activated, causing the release of atrial natriuretic peptide (also known as ANP) from the right atrium of the heart. Together, these actions cause the nephrons to keep more sodium and water in the filtrate, which depletes the volume of extracellular fluid and moves blood pressure closer to the baseline. When blood pressure is low, kidneys secrete renin, an enzyme that sets off a series of physiological reactions that result in the tubules reabsorbing more water and sodium as well as vasoconstriction in blood vessels. Together, these activities raise blood pressure back to the baseline. The nephrons are also induced to increase water retention by the secretion of vasopressin (also known as antidiuretic hormone, ADH), which is secreted by the hypothalamus in response to feedback from baroreceptors (pressure sensors) in the aortic arch and carotid sinus.

16.6 Summary

Kidneys as well as gills, integument, and various glands are all important for maintaining both water balance and ion balance in the vertebrate body. Vertebrates show a variety of adaptations and variations of these organs to meet the specific challenges of their particular environments.

16.7 Further Reading

Check out the hashtag #nephmadness online. Not only did it inspire some of the fun, unusual examples used in this chapter; there are a lot of amazing human renal physiology facts that you can learn about!

You can read more about Dr. Jonathan Resiman’s love of urine (and anatomy in general) in his book The Unseen Body, 2021, Flatiron Press.

16.8 References

1. Baitchman, Eric J., and George V. Kollias. “Clinical anatomy of the North American river otter (Lontra canadensis).” Journal of Zoo and Wildlife Medicine 31 (2000): 473–483.
2. de Bakker, B. S., M. J. B. van den Hoff, P. D. Vize, and R. J. Oostra. “The pronephros; a fresh perspective.” Integrative and Comparative Biology 59 (2019): 29–47.
3. Baldwin, G. F., and P. J. Bentley. “Roles of the skin and gills in sodium and water exchanges in neotenic urodele amphibians.” American Journal of Physiology: Regulatory, Integrative, and Comparative Physiology 242 (1982): R94–R96.
4. Braun, Marvin H., and Steve F. Perry. “Ammonia and urea excretion in the Pacific hagfish Eptatretus stoutii: Evidence for the involvement of Rh and UT proteins.” Comparative Biochemistry and Physiology Part A 157 (2010): 405–415.
5. Chambers, Joseph M., and Rebecca A. Wingert. 2020. “Advances in understanding vertebrate nephrogenesis.” Tissue Barriers 8 (2020): 1832844.
6. Dantzler, William G. Comparative Physiology of the Vertebrate Kidney, 2nd ed. New York: Springer Nature, 2016.
7. Davis, Jon R., and Dale F. DeNardo. “The urinary bladder as a physiological reservoir that moderates dehydration in a large desert lizard, the Gila monster Heloderma suspectum.” Journal of Experimental Biology 210 (2007): 1472–1480.
8. Ditrich, Hans. “The origin of vertebrates: A hypothesis based on kidney development.” Zoological Journal of the Linnean Society 150 (2007): 435–441.
9. Evans, David H., Peter M. Piermarini, and Keith P. Choe. “Homeostasis: Osmoregulation, pH regulation, and nitrogen excretion.” In Biology of Sharks and Their Relatives, Vol. 1, Ed 1., edited by Jeff C. Carrier, Jack A. Musick, and Michael R. Heithaus, 247–268. Boca Raton: CRC Press, 2004.
10. Evans, Roger G. “Evolution of the glomerulus in a marine environment and its implications for renal function in terrestrial vertebrates.” American Journal of Physiology: Regulatory, Integrative, and Comparative Physiology 324 (2023): R143–R151.
11. Fels, Lüder M., Sabine Kastner and Hilmar Stolte. “The hagfish kidney as a model to study renal physiology and toxicology.” In: The Biology of Hagfishes, edited by Jørgen Mørup Jørgensen, Jens Peter Lomholt, Roy E. Weber, and Hans Malte, 347–363. London: Chapman and Hall, 1998.
12. Ferreira-Martins, D., J. M. Wilson, S. P. Kelly, D. Kolsov, and S. D. McCormick. “A review of osmoregulation in lamprey.” Journal of Great Lakes Research 47 (2021): S59–S71.
13. Finlay, Ross W., Russell Poole, Ger Rogan, Eileen Dillane, Deirdre Cotter, and Thomas E. Reed. “Hyper- and hypo-osmoregulatory performance of Atlantic salmon (Salmo salar) Smolts infected with Pomphorhynchus tereticollis (Acanthocephala).” Frontiers in Ecology and Evolution 9 (2021): 689233.
14. Geven, Edwin J. W. and Peter H. M. Klaern. “The teleost head kidney: Integrating thyroid and immune signaling.” Developmental Comparative Immunology 66 (2017): 73–83.
15. Hazard, Lisa C. “Ion secretion by salt glands of desert iguanas (Dipsosaurus dorsalis).” Physiological and Biochemical Zoology: Ecological and Evolutionary Approaches 74 (2001): 22–31.
16. Heimer, Rachel A., Belinda J. Davis, and John A. Donald. “The effect of water deprivation on the expression of atrial natriuretic peptide and its receptors in the spinifex hopping mouse, Notomys alexis.” Comparative Biochemistry and Physiology Part A: Molecular and Integrative Physiology 132 (2002): 893–903.
17. Hyodo, Susumu, Keigo Kakumura, Wataru Takagi, Kumi Hasegawa, and Yoko Yamaguchi. “Morphological and functional characteristics of the kidney of cartilaginous fishes: With special reference to urea reabsorption.” Regulatory, Integrative and Comparative Physiology 307 (2014): R1381–R1395.
18. Khan, Kanawar Nasir M., Gordon C Hard, and Carl L. Alden. “Kidney.” In Haschek and Rousseaux’s Handbook of Toxicologic Pathology, Vol. 3, 3rd ed., edited by Wanda M. Haschek, Colin G. Rousseaux and Matthew A. Wallig, 1667–1773. Cambridge: Academic Press, 2013.
19. Larsen, Erik Hviid, Lewis E. Deaton, Horst Onken, Michael O’Donnell, Martin Grosell, William H. Dantzler, and Dirk Weihrauch. “Osmoregulation and excretion.” Comparative Physiology 4 (2014): 405–573.
20. Maluf, N. S., and J. J. Gassman. “Kidneys of the killer whale and significance of reniculism.” Anatomical Record 250 (1998): 34–44.
21. Marshall, W. S. “Independent Na⁺ and Cl^– active transport by urinary bladder of epithelium of brook trout.” American Journal of Physiology 250 (1986): R227–R234.
22. Martin Jon A., and Stan S. Hillman. “The physical movement of urine from the kidneys to the urinary bladder and bladder compliance in two anurans.” Physiological and Biochemical Zoology: Ecological and Evolutionary Approaches 82 (2009): 163–169.
23. Møbjerg, N., E. H. Larsen, and Å. Jespersen. “Morphology of the kidney in the larvae of Bufo viridis (Amphibia, Anura, Bufonidae).” Journal of Morphology 245 (2000): 177–195.
24. Oh Han Young, So Ryung Shin, Jung Jun Park, Hyeon Jin Kim, and Jung Sick Lee. “Distribution of nephrons in the head kidney of three species of Sebastes (Teleostei: Scorpaenidae).” Journal of Fish Biology. 103 (2023): 965–973.
25. Ojeda, José L., Wai P. Wong, Yuen K. Ip, and José M. Icardo. “Renal corpuscle of the African lungfish Protopterus dolloi: Structural and histochemical modifications during aestivation.” Anatomical Record 291 (2008): 1156–1172.
26. OpenStax College. “Hormonal Control of Osmoregulatory Functions,” 2012. https://pressbooks-dev.oer.hawaii.edu/biology/chapter/hormonal-control-of-osmoregulatory-functions/.
27. Pannabecker, Thomas L. “Aquaporins in desert rodent physiology.” Biological Bulletin 229 (2014): 120128.
28. Pillans, Richard D., and Craig E. Franklin. “Plasma osmolyte concentrations and rectal gland mass of bull sharks Carcharhinus leucas, captured along a salinity gradient.” Comparative Biochemistry and Physiology Part A 138 (2004): 363–371.
29. Resendale, Albina D., Alexandre Lobo-da-Cunha, Fernanda Malhão, Filipa Franquinho, Rogério A F Monteiro, and Eduardo Rocha. “Histological and stereological characterization of Brown Trout (Salmo trutta f. fario) trunk kidney.” Microscopy and Microanalysis 16 (2010): 677–687.
30. Roberts, Ronald J. Fish Pathology, 4th ed. Hoboken: John Wiley & Sons, 2012.
31. Sakai, Tatsuo, Ralph Billo, and Wilhelm Kriz. “The structural organization of the kidney of Typhlonectes compressicadus (Amphibia, Gymnophiona).” Anatomy and Embryology 174 (1986): 243–252.
32. Senerat, Sinlapachai, Jes Kettratad, Supanut Pairohakul, Sumate Ampawong, Brian P. Huggins, Melissa M. Coleman, and Gen Kaneko. “An update on the evolutionary origin of aglomerular kidney with structural and ultrastructural descriptions of the kidney in three fish species.” Journal of Fish Biology 100 (2022): 1283–1298.
33. Sielgel, Dustin S., David M. Sever, and Robert Aldrige. “The pelvic kidney of male Ambystoma maculatum (Amphibia, Urodela, Ambystomatidae) with special reference to the sexual collecting ducts” Journal of Morphology 271 (2010): 1422–1439.
34. Takei, Yoshio, Ray C. Bartolo, Hiroaki Fujihara, Yoichi Ueta and John A. Donald. “Water deprivation induces appetite and alters metabolic strategy in Notomys alexis: Unique mechanisms for water production in the desert.” Proceedings of the Royal Society B 279 (2012): 2599–2608.
35. Velasquez, Maneul T., Ali Ramezani, Alottaibi Manal, and Dominic S. Raj. “Trimethylamine N-oxide: The good, the bad and the unknown.” Toxins 8 (2016): 326.
36. Wadei, Hani M., and Stephen C. Textor. “The role of the kidney in regulating arterial blood pressure.” Nature Reviews Nephrology 8 (2012): 602–609.
37. Wake, Marvalee H. “Evolutionary morphology of the caecilian urogenital system: II. The kidneys and urogenital ducts.” Acta Anatomica 75 (1970): 321–358.
38. Wake, Marvalee H. “Urogenital morphology of dipnoans, with comparisons to other fishes and amphibians.” Journal of Morphology, 190, Suppl. 1 (1986): 199–216.
39. Weiner, Scott V. “Effects of the environment on the evolution of the vertebrate urinary tract.” Nature Reviews Urology 20 (2023): 719–738.
40. Zhou, Xu, Wenqi Rong, Boxiong Guo, Xiaofang He, Li Cao, Yu Zheng, Shixia Xu, and Guang Yang. “The evolution of the discrete multireniculate kidney in mammals from ecological and morphological perspectives.” Genome Biology and Evolution 15 (2023): evad075.