Regenerative Nephrology

By Michael S. Goligorsky

Since the publication of the first edition of this book in 2010, an explosion of spectacular discoveries in the field of regeneration has compelled the current revisit of the field of Regenerative Nephrology. This second edition features subjects as diverse as age and gender influencing regenerative processes; mechanisms and pathways of premature cell senescence affecting kidney regeneration; the ways intrinsic regenerative processes can become subverted by noxious stressors eventuating in disease progression; novel mechanistic and engineering efforts to recreate functional kidney or its component parts; cell reprogramming and reconditioning as emerging tools of future regenerative efforts; and effects of various biologicals on kidney regeneration. These newer additions to the armamentarium of Regenerative Medicine and Nephrology have become an integral part of the second edition of the book. Cutting-edge investigations are summarized by the constellation of the most experienced contributing authors coming together from around the world under the umbrella of the second edition.

• A significant expansion of section on induced pluripotent cells and trajectories of their differentiation. This will be followed by mechanisms and modalities of cell reprogramming for therapeutic purposes
• A new section on tissue engineering of the kidney of interest to nephrologists and urologists
• An entire section dedicated to causes of regenerative failure with the emphasis on recent discoveries of senescent cells in kidney disease, pathologic effects of senescent cells, advents in senotherapies and rejuvenation therapies
• A vastly expanded section on pharmacotherapies promoting kidney regeneration, trials of engineered organs, manufacturing in regenerative medicine and smooth transition to the clinical trials, with an update on some ethical issues

• Language: English
• Publisher: Academic Press
• Release date: Jun 12, 2021
• ISBN: 9780128233191

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Part I

Kidney development and regeneration

Chapter 1: Nephrogenesis in health and disease

Adrian S. Woolfa,b; Sophie L. Ashleya,c    a Division of Cell Matrix Biology and Regenerative Medicine, School of Biological Sciences, Faculty of Biology Medicine and Health, University of Manchester, Manchester, United Kingdom

b Royal Manchester Childreńs Hospital, Manchester University NHS Foundation Trust, Manchester Academic Health Science Centre, Manchester, United Kingdom

c Manchester Medical School, University of Manchester, Manchester, United Kingdom

Abstract

The mammalian renal tract is a physically integrated group of organs whose main functions are to generate and void urine. The renal tract comprises the kidney and the urinary tract, the latter incorporating the renal pelvis, the ureter, the urinary bladder, and the urethra. The embryonic origin and subsequent shaping, or morphogenesis, and cellular differentiation of the renal tract are multistep processes, so they are unsurprisingly prone to error. This chapter addresses the normal development of the renal tract, with an emphasis on the early steps of development of the metanephric kidney. We then proceed to consider certain genetic and environmental causes of renal tract malformations. Given that the theme of this book is regeneration, we consider whether aberrations of renal tract development can be rescued by manipulation of, for example, growth factor signaling.

Keywords

Agenesis; Collecting duct; Dysplasia; Environment; Gene; Hypoplasia; Malformation; Nephron; Ureter; Ureteric bud

Acknowledgments

We acknowledge grant support as follows: the Medical Research Council (MR/K026739/1 and MR/T016809/1): Kidney Research UK (Intercalating Student Award); and Kidneys for Life (Pump Priming project grant 2018 to ASW).

Conflict of interest statement

Neither author has conflicts of interest to declare.

Introduction

The mammalian renal tract is a physically integrated group of organs whose main functions are to generate and void urine [1, 2]. The renal tract comprises the kidney and the urinary tract, the latter incorporating the renal pelvis, the ureter, the urinary bladder, and the urethra. The embryonic origin and subsequent shaping, or morphogenesis, and cellular differentiation of the renal tract are multistep processes, so they are unsurprisingly prone to error. This chapter addresses the normal development of the renal tract, with an emphasis on the early steps of development of the metanephric kidney. We then proceed to consider some genetic and environmental causes of renal tract malformations. Given that the theme of this book is regeneration, we consider whether aberrations of renal tract development can be rescued by manipulation of, for example, growth factor signaling.

Experimental models

The mechanisms of mammalian renal tract development have been most extensively studied in rodents, although the anatomy of human renal tract development has been described in detail by Edith Potter over 50 years ago [3]. The anatomical steps of renal tract development are similar in mice, rats, and humans, although the timetable is accelerated in the rodents. There are also differences of scale between humans and rodents. For example, the healthy human kidney contains around 1.4 million glomeruli [4], while a healthy mouse [5] or rat [6] kidney respectively contain around 6 and 30 thousand glomeruli. Our focus is mammalian development but readers who are interested in how kidneys form in frogs, fish, and flies are directed to comprehensive reviews published elsewhere [1]. Although covered in detail in other chapters, we briefly allude to pluripotent stem cell technology which is increasingly being used to study kidney development, complementing the study of native renal tracts.

Developmental events preceding kidney development

The primitive streak arises toward the posterior end of the back of the early embryo. The streak coordinates the migration of cells from the surface of the embryo such that they become the three germ layers: the endoderm, mesoderm, and ectoderm. These gastrulation events are driven by gradients of growth factors such as wingless-related integration site-3A (WNT3A) and bone morphogenetic protein 4 (BMP4) [7, 8]. The portion of mesoderm called the intermediate mesoderm will give rise to the embryonic kidneys, of which there are three sets of paired organs. The pronephric and mesonephric kidneys form but then regress in the embryonic period and they are not discussed further here. In contrast, the metanephric kidney, or metanephros, forms and survives to become the definitive mammalian kidney. It initiates on day 10 of mouse gestation, day 12 of rat gestation, and week 5 of human gestation.

Cell populations in the metanephric kidney

The metanephros contains two populations of precursor cells, the ureteric bud and the metanephric mesenchyme, as depicted in Fig. 1.1 [9]. Based on mouse experiments [10], ureteric bud precursor cells express the transcription factor called odd-skipped-related 1 (OSR1). They are more anteriorly positioned within the intermediate mesoderm than the precursor cells that will become metanephric mesenchyme; the latter express the T-box transcription factor T (TBXT, or brachyury). The ureteric bud is an epithelial tube that branches from the caudal part of the Wolffian duct of which there are two, each formed from the intermediate mesoderm and extending caudally toward the cloaca, the precursor of the urinary bladder and hindgut [11]. The stalk of the ureteric bud will give rise to the urothelium of the ureter, while its top will branch serially to form the collecting ducts of the metanephric kidney. The first sets of branches are remodeled to become the epithelia of the renal pelvis [2]. Therefore, all these derived cells are said to be in the ureteric bud lineage. The metanephric mesenchyme will form the nephrons (i.e., podocytes, proximal tubule, loop of Henle, and distal convoluted tubule), and additionally contributes to kidney interstitial cells. Moreover, a subset of cells in the metanephric mesenchyme express markers of endothelial precursors and, at least experimentally, can contribute to kidney interstitial and glomerular capillaries [12]. Fig. 1.2 shows histology of three time points from the rat metanephros where early ureteric bud branch tips and primitive nephrons are evident.

Fig. 1.1

Fig. 1.1 Cell lineages in the embryonic metanephros. The frame on the left shows the histology of the metanephros at its inception, with a central ureteric bud surrounded by metanephric mesenchyme. Bar = 50 μm. The frame on the right depicts mutual induction between these compartments. The ureteric bud differentiates into the urothelial stalk of the ureter and the arborizing collecting ducts within the kidney. The metanephric mesenchyme undergoes mesenchymal to epithelial transition to form nephrons, comprising glomerular and tubule epithelia, whereas other cells in the mesenchymal compartment will form interstitial cells and endothelia. Reproduced from Woolf AS. Growing a new human kidney. Kidney Int 2019;96:871–882.

Fig. 1.2

Fig. 1.2 Bright field images of maturation of the embryonic kidney of the rat. Sections from control animals were stained with hematoxylin and eosin. (A and B) Embryonic day 13 metanephros with ureteric bud branch tips (u) and condensing mesenchyme (m). Nearby vessels are indicated (v). (C and D) Embryonic day 15 metanephros shows considerable growth and differentiation but the most mature nephron stages are S-shaped bodies (s) and an outer nephrogenic zone is still prominent. (E and F) Two-week postnatal kidney. Note that there is no nephrogenic zone and that all glomeruli (g) appear mature with capillary loops. Bars are: 100 μm in A, C and E, and 10 μm in B, D and F. Reproduced from Welham SJM, Wade A, Woolf AS. Protein restriction in pregnancy is associated with increased apoptosis of mesenchymal cells at the start of rat metanephrogenesis. Kidney Int 2002;61:1231–1242.

Molecular control of kidney development: The ureteric bud/collecting duct lineage

As reviewed [13], the correct point of emergence of the ureteric bud is controlled by a balance of stimulatory and inhibitory growth factors. For example, BMP4, secreted by mesoderm that is not fated to become metanephric mesenchyme, inhibits emergence, whereas glial cell line-derived neurotrophic factor (GDNF) and fibroblast growth factor 10 (FGF10), both secreted by the definitive metanephric mesenchyme, stimulate bud emergence. Within the domain of the Wolffian duct where the bud will emerge, cells upregulate rearranged during transfection (RET), the receptor tyrosine kinase that binds GDNF [14]. The Fraser extracellular matrix complex subunit 1 (FRAS1) protein, and family members FRAS1-related extracellular matrix 1 (FREM1) and FREM2, together coat the surface of the ureteric bud and are required to optimize GNDF signaling [15, 16]. The FRAS1/FREM1/FREM2 complex also facilitates the binding of the matrix molecule nephronectin to integrin α8β1 that is required for physical interaction between the ureteric bud and metanephric mesenchyme [17]. Fine-tuning of branching is mediated by an intrinsic ureteric bud molecule called sprouty-1 which antagonizes GDNF and FGF10 signaling. Sprouty-1 itself is downregulated by angiotensin II [18], and BMP4 is antagonized by yet another secreted factor, gremlin-1 [19]. Serial branching of the ureteric bud forms the collecting duct tree, a process again primarily driven by GDNF, secreted by the cortical rim of undifferentiated mesenchyme. The formation of the tree is fine-tuned by numerous other molecules including those encoded by the neurofibromatosis 2 gene NF2 [20] and the planar cell polarity genes called Van Gogh-like protein 2 (VANGL2) and cadherin EGF LAG seven-pass G-type receptor 1 (CELSR1) (Fig. 1.3) [21].

Fig. 1.3

Fig. 1.3 Patterns of ureteric bud branching in mice with mutations in planar cell polarity genes. Images of whole metanephroi from embryonic day 13 wild-type (A), Celsr1 Crsh/+ (C), Celsr1 Crsh/Crsh (E), Vangl2 Lp/+ (G), and Celsr1 Crsh/+ : Vangl2 Lp/+ (I) mice stained with calbindin-D28K and visualized by optical projection tomography. Branching networks of individual metanephroi from embryonic day wild-type (B), Celsr1 Crsh/+ (D), Celsr1 Crsh/Crsh (F), Vangl2 Lp/+ (H), and Celsr1 Crsh/+ : Vangl2 Lp/+ (J) mice. Note the impaired branching patterns in the homozygous mutant and compound heterozygous mutant embryonic kidneys. Reproduced from Brzóska HL, d’Esposito AM, Kolatsi-Joannou M, Patel V, Igarashi P, Lei Y, et al. Planar cell polarity genes Celsr1 and Vangl2 are necessary for mammalian kidney growth, differentiation and rostrocaudal patterning. Kidney Int 2016;90:1274–1284.

Molecular control of kidney development: The metanephric mesenchyme/nephron lineage

Metanephric mesenchymal cells express the Wilms tumor 1 (WT1) transcription factor, without which they regress prematurely [22], another aberrant pathway to renal agenesis. In health, cells in this compartment undergo rounds of mesenchymal-to-epithelial transition to form multiple layers of nephrons (Fig. 1.2), the deepest layers having formed first. In mice, the waves of nephron formation, literally nephrogenesis, continue until a week after birth, whereas in humans the process is complete by 34 weeks of gestation. At the same time as layers of nephrons are being generated, the expression of FGF9 and FGF20 growth factors and the sine oculis homeobox homolog 2 (SIX2) transcription factor preserve a rim of metanephric mesenchyme that will give rise to further rounds of nephrons; without these growth molecules, there is premature exhaustion of nephron progenitors [23, 24]. During the development of the metanephric kidney, subsets of cells have a high proliferative rate, especially those in the branching tips of the ureteric tree, and in primitive nephrons [25]. Concurrently, apoptctic death takes place in the developing metanephros, and it has been estimated that around half the cells generated in the metanephros will undergo programmed cell death during development [26]. Apoptosis is prominent in cells around emerging nephrons, as depicted in Fig. 1.4 [27]. Here, perhaps interstitial cells need to be deleted to facilitate sculpting of the nephron. Apoptosis is also prominent in the deep medulla, perhaps to make way for loops of Henle that grow into the forming papilla. Indeed, if apoptosis is experimentally inhibited in metanephric organ culture, then nephrogenesis is perturbed [28].

Fig. 1.4

Fig. 1.4 Confocal laser scanning images of the embryonic day 13 rat metanephros. Sections from control diets. (A and B) The same area under appropriate wavelengths for detection of terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) and propidium iodide-labeled nuclei, respectively. Two TUNEL-labeled nuclei ( bright green ) associated with condensing mesenchyme are indicated in panel A ( arrows ) and appear as pyknotic nuclei stained red with propidium iodide ( arrows in panel B). A ureteric bud branch (u) and mesenchyme (m) are indicated. The arrowhead indicates a mitotic cell. (C) Whole embryonic day metanephros, outlined for clarity by a dotted line , and surrounding tissues. The photograph was generated by merging the signals from both wavelengths, with apoptotic TUNEL/propidium iodide-stained nuclei appearing bright yellow . Note the apoptotic cells, often in clusters in loose mesenchyme (*). Apoptosis was also seen, but less prominently, in condensed mesenchyme (cm) and was not detected in ureteric bud branches (u). Bars are 10 μm in A and B, and 30 μm in C. Reproduced from Welham SJM, Wade A, Woolf AS. Protein restriction in pregnancy is associated with increased apoptosis of mesenchymal cells at the start of rat metanephrogenesis. Kidney Int 2002;61:1231–1242.

The morphological journey to become a nephron begins close to a ureteric bud branch tip, or ampulla, where a subset of cells in the metanephric mesenchyme are induced to aggregate and then form a vesicle, essentially an epithelial sphere. The vesicle elongates and curves via stages called the comma and S-shaped body. During elongation, the maturing nephron patterns, proximally–distally, into glomerular podocytes, the proximal tubule, the loop of Henle, and the distal convoluted tubule. The latter fuses with a collecting duct to form a patent conduit. WNT4 is a key molecule that drives the generation of primitive nephron vesicles from mesenchymal precursors [29]. The transmembrane molecule called neurogenic locus notch homolog protein 2 (NOTCH2) is required for differentiation of the proximal segment of the primitive nephron segments (i.e., podocytes and the proximal tubule) [30]. These proximal segment fates are also controlled by hepatocyte nuclear factor-1B (HNF1B) [31], a transcription factor that also has roles in controlling mitochondrial respiration in kidney tubules [32] and enhancing the differentiation of collecting duct epithelia [33].

Glomerular podocytes differentiate from cuboidal into mature flattened epithelia that extend foot processes from their basal surface which abut on the glomerular basement membrane [34]. The maturing podocytes upregulate WT1 to levels higher than that found in metanephric mesenchyme [25]. This is accompanied by the expression of genes that code for slit diaphragm proteins, including nephrin and podocin. At the same time, the composition of the glomerular basement membrane matures to include collagen IV α3, α4, and α5, as well as laminin B2. Of note, this protein maturation fails to take place in organ culture, a scenario where the glomerular tuft lacks capillary loops that are found in vivo. In native developing kidneys, maturing podocytes secrete vascular growth factor A and platelet-derived growth factor B, and gradients of these molecules attract precursor cells to migrate into the tuft and respectively form capillary loops [35] and mesangial cells [36]. Of note, vascularized glomeruli expressing basement membrane can be generated after subcutaneous implantation of human pluripotent stem cell-derived kidney precursor cells into immunocompromised mice [37], as depicted in Fig. 1.5.

Fig. 1.5

Fig. 1.5 Mature glomeruli formed in vivo after implanting human pluripotent stem cell-derived kidney precursor cells into immunocompromised mice. Glomerular tufts immunostain ( brown ) for the podocyte marker synaptopodin. Glomeruli immunostain for collagen α3 (IV) and vascular endothelial growth factor A (VEGFA). Capillaries that immunostain for platelet endothelial cell adhesion molecule (PECAM) are detected inside the glomerular tuft. Nuclei in the collagen α3 (IV) and PECAM frames are counterstained with hematoxylin. Bar = 50 μm. Reproduced from Woolf AS. Growing a new human kidney. Kidney Int 2019;96:871–882; Adapted from Bantounas I, Ranjzad P, Tengku F, Silajdžić E, Forster D, Asselin MC, et al. Generation of functioning nephrons by implanting human pluripotent stem cell-derived kidney progenitors. Stem Cell Reports 2018;10:766–779.

Urinary tract development

The urinary tract comprises the renal pelvis, the ureter, the urinary bladder, and the urethra. The ureters propel urine from the renal pelvis to the bladder. In turn, the bladder stores urine at low pressure until the detrusor smooth muscle in the bladder wall contracts, resulting in complete voiding of its contents through the urethra. The mammalian ureter, a wholly mesodermal-derived organ, initiates when the ureteric bud branches from the Wolffian duct. The epithelial bud then elongates to form the stalk of the ureter, at the same time differentiating to form the pseudostratified urothelium of the ureter and renal pelvis [38]. The urothelium expresses uroplakin proteins that confer a water-tight barrier, preventing the egress of urine out of the urinary tract [39]. Concurrently, mesenchyme condenses around the urothelial tube and differentiates into a smooth muscle [40]. The urothelial layer of the bladder differentiates from the endoderm-derived urogenital sinus, and adjacent mesoderm-derived mesenchymal cells differentiate into smooth muscle. The embryonic urothelium acts as a signaling center [41, 42], secreting sonic hedgehog (SHH) that acts on nearby mesenchymal cells to drive their proliferation and differentiation into smooth muscle. This involves the muscle precursor cells secreting BMP4 that leads to the transcription factors teashirt-3 and myocardin upregulating the expression of smooth muscle contractile proteins such as smooth muscle actin and myosin heavy chain [40, 43]. The caudal end of the Wolffian duct fuses with the urogenital sinus, facilitated by RET [44], and then involutes through vitamin A-mediated apoptosis [45]; the result is that the distal end of the forming ureter becomes incorporated into the lateral aspect of the bladder where the uretero-vesical junction acts as a valve that prevents reflux of urine. The bladder is richly innervated [46] and, in humans, from the late first trimester autonomic nerves are visualized, extending between smooth muscle bundles [47]. These postganglionic neurons emanate from autonomic ganglia outside the bladder and they express heparanase 2 (HPSE2) and leucine-rich repeats and immunoglobulin-like domains protein 2 (LRIG2) proteins that appear to pattern their distribution [48]. Urinary tract differentiation harmonizes with kidney development, so that the tract is ready to receive and transmit urine made by the developing kidney. Indeed by the end of the first trimester of human gestation, the bladder is a muscular organ that receives and voids urine.

As yet, protocols have not been published that describe the simultaneous generation of both a kidney and ureter from pluripotent stem cells. However, such stem cells can be driven toward a urothelial fate and, when recombined with native mouse intermediate mesoderm from near the Wolffian duct, a ureter-like contractile tube can be formed [49].

As depicted in Fig. 1.6, a chimeric renal organoid containing nephrons derived from human pluripotent stem cells enveloping an embryonic mouse ureter can be generated by tissue engineering [50].

Fig. 1.6

Fig. 1.6 A chimeric organoid containing nephrons derived from human pluripotent stem cells and an embryonic mouse ureter. (A) The organoid visualized in culture with the mouse ureter (the vertical tube inside the red box) surrounded by human nephron-like tissues. (B) Histology of the boxed zone shows the mouse ureter (u) flanked by glomeruli (g) in this section stained with hematoxylin and eosin. (C) Nearby section immunostained with an antibody to human mitochondria ( brown positive signal ) confirms that the glomeruli are of human origin. (D) The embryonic mouse ureter has differentiated to form an α-smooth muscle actin positive (α-SMA) coat ( brown ), as it does in vivo. Bar in A is 500 μm, and bars in B–D are 50 μm. Adapted from Ashley S. Plumbing kidney organoids derived from human pluripotent stem cells. M Res Thesis, University of Manchester. 2019, pp. 1–100.

Abnormal development of the renal tract

The embryonic origin and subsequent shaping, or morphogenesis, and cellular differentiation of the renal tract are multistep processes, so they are unsurprisingly prone to error. Renal tract malformations, sometimes also called Congenital Anomalies of the Kidney and Urinary Tract (CAKUT), are a collection of rare disorders, yet this disease category is the primary diagnosis in half of all children with end-stage renal disease (ESRD) [51]. Such malformations also account for up to a fifth of young adults with ESRD [52]. Moreover, a third of all anatomical defects detected by prenatal ultrasound scan screening are renal tract malformations, and the most severely affected fetuses often undergo elective termination of pregnancy [53]. Increasingly, human renal tract malformations are being found to have genetic bases [54, 55], although it is likely that environmental perturbations also can occur.

Models of renal agenesis

The emergence of the ureteric bud from the Wolffian duct can be considered as the critical event in the formation of the metanephric kidney. Indeed, if the bud does not initiate and grow into the metanephric mesenchyme, the result is an absent kidney and ureter, or renal agenesis. Renal tract development will also be compromised if the bud emerges either too high (cranial) or too low (caudal) along the Wolffian duct. These respective scenarios will result in a malformed (dysplastic) kidney or a ureter with a too lateral insertion into the bladder. A third possible aberration is the emergence of multiple ureteric buds, leading to a duplicated (duplex) renal tract. An example of renal agenesis is provided by Fraser syndrome where individuals carry biallelic mutations of either FRAS1 or FREM2 and often lack both kidneys [15, 16]. As discussed earlier, the encoded proteins coat the surface of the ureteric bud and facilitate paracrine signaling by mesenchymal-derived growth factors. Remarkably, this most severe of all renal tract malformation is not written in stone, at least in an experimental mouse model. Examination of Fras1 homozygous mutant mice shows that the ureteric bud either fails to initiate or, if it does, it fails to fully engage with the metanephric mesenchyme. The result in vivo is that the mesenchyme dies and later in gestation kidney tissue cannot be found. If the zone comprising the mutant caudal Wolffian duct and surrounding intermediate mesoderm is explanted in organ culture, a similar sequence of events occurs [15]. If, however, the culture is supplemented with high concentrations of GDNF or FGF10, the metanephric development proceeds such that a small rudiment is formed. Strikingly, if the homozygous Fras1 null mutation is placed on a genetic background where sprouty-1 expression is less than normal, kidney development proceeds so that two kidneys are generated that can sustain the life of Fras1 mutant through to adulthood [16].

There are also environmental causes of renal agenesis. For example, overexposure of developing embryonic mice, before the kidney initiates, to retinoic acid, a metabolite of vitamin A, leads to the downregulation of WT1 in metanephric mesenchyme which proceeds to degenerate with upregulated apoptosis [56]. Again, as for the Fras1 genetic example above, this sequence leading to agenesis can be mimicked in vitro when rudiments, harvested from retinoic acid exposed mice, are explanted in organ culture (Fig. 1.7). Notably, the development of these rudiments can be rescued by the addition of high concentrations of serum [56]. Perhaps a yet-to-be defined growth factor present in serum mediates this effect, and it has been shown that the addition of epidermal growth factor can minimize apoptosis in otherwise normal embryonic rodent kidneys [26].

Fig. 1.7

Fig. 1.7 Mouse metanephric explants cultured in vitro for 5 days. (A–D) Morphology of explanted metanephroi from control (CT) or retinoic acid-treated (RA) embryos cultured in serum-free medium (A and B), or medium supplemented with 10% (C) or 20% serum (D). E and F. Histology of RA-treated metanephric explants cultured in 20% serum showed that the rudiments had undergone differentiation, with formation of condensates, primitive tubules, and avascular glomeruli ( arrowhead ). (G) Calbindin immunostaining of UB branch epithelium showed multiple branching in metanephric explants from RA-treated embryos after culture in 20% serum. (H) Real-time quantitative RT-PCR analyses of expression levels of Wt1 relative to β- actin in metanephroi from control and RA-exposed embryos at 2 days after culture in serum-free medium or medium supplemented with 20% serum [*, P  < 0.0001, CT versus RA in the same culture condition; † , P  < 0.005, CT (no serum) versus CT (20% serum); ‡ , P  < 0.0001, RA (no serum) versus RA (20% serum); Student’s t -test]. Scale bar: 0.5 mm (A–D, G), 0.35 mm (E), 1.4 mm (F). Reproduced from Tse HK, Leung MB, Woolf AS, Menke AL, Hastie ND, Gosling JA, et al. Implication of Wt1 in the pathogenesis of nephrogenic failure in a mouse model of retinoic acid-induced caudal regression syndrome. Am J Pathol 2005;166:1295–1307.

Models of renal dysplasia

Another phenotype of human kidney maldevelopment is renal dysplasia, in which the kidney begins to form but its differentiation is incomplete, with impaired branching morphogenesis and nephrogenesis, and metaplastic, for example, with the generation of smooth muscle and cartilage (Fig. 1.8). As for renal agenesis, there are genetic and environmental causes. An example of the former are heterozygous mutations of HNF1B, currently the commonest genetically defined cause of human renal dysplasia [54, 57, 58]. As discussed earlier, this gene codes for a transcription factor with multiple roles in morphogenesis and differentiation of epithelia in both the ureteric bud and metanephric mesenchymal lineages. Human renal dysplastic kidneys have been shown to display a significantly increased number of apoptotic cells compared with time-matched healthy control kidneys [59], perhaps explaining the tendency of these organs to regress in size either before birth or in the first years of childhood.

Fig. 1.8

Fig. 1.8 The histology of human renal dysplasia stained with hematoxylin and eosin. (A) Dysplastic tubule (*marks the lumen) surrounded by poorly differentiated stromal tissues(s). (B) Metaplastic cartilage (c). Note the absence of normal nephrons and glomeruli. Bars are 15 μ in A and 150 μ in B. Reproduced from Bingham C, Ellard S, Cole TR, Jones KE, Allen LI, Goodship JA, et al. Solitary functioning kidney and diverse genital tract malformations associated with HNF1b mutations. Kidney Int 2002;61:1243–1251.

Renal dysplasia, however, is also frequently observed in association with fetal urinary flow impairment [43]. Indeed, impaired nephrogenesis can be experimentally generated by, for example, ligating the ureter of a fetal sheep [60], as depicted in Fig. 1.9. Similar disruption of nephrogenesis follows experimental ligation of the fetal urethra [61]. This physical insult triggers upregulation of transforming growth factor β (TGFβ) in the kidney [60]. Moreover, the addition of this factor to an epithelial cell line derived from a human dysplastic kidney leads to transdifferentiation to a smooth-muscle-like phenotype, as shown in Fig. 1.10 [62]. The addition of TGFβ to mouse embryonic ureteric explants retards their linear growth; this is associated with a tendency for downregulation of transcripts encoding FGF10, and exogenous FGF10 can partially rescue the linear growth defect in TGFβ-exposed ureteric explants [38].

Fig. 1.9

Fig. 1.9 Ureteric ligation leads to impaired kidney development and upregulation of TGFB1 expression. TGFβ1 in situ hybridization in sham-operated and obstructed sheep fetal kidneys. A and C are sections of sham-operated developing kidneys, whereas B, D, and E are from obstructed kidneys. All panels represent the results of hybridization with TGFβ1 antisense probes, apart from E in which a control sense probe was used. (A) TGFβ1 transcripts were detected in ureteric bud (u) and forming nephrons (n) in the normal superficial cortex. (B) Prominent signal for TGFβ1 was observed in dilated tubule epithelia ( arrowheads ) and glomerular tufts (t; arrows ) in the obstructed cortex. (C) Faint TGFβ1 signal was detected in sham-operated developing medullary collecting ducts ( arrows ). (D) In contrast, there was TGF-β1 transcript up-regulation in the larger dilated collecting ducts (cd) in the obstructed kidneys. (E) Only background signal was detected using the sense probe. Scale bar, 15 μm. Reproduced from Yang SP, Woolf AS, Quinn F, Winyard PJD. Deregulation of renal transforming growth factor-β1 after experimental short-term ureteric obstruction in fetal sheep. Am J Pathol 2001;159:109–117.

Fig. 1.10

Fig. 1.10 Effects of TGFβ1 on morphology and tight junctions in an epithelial cell line derived from a human dysplastic kidney. Gross morphology is shown in A and B, whereas C and D show tight junction zonula occludens-1 (ZO1) immunocytochemistry (with propidium-iodide nuclear counterstaining) of epithelial-like dysplastic cells cultured in either control medium (A and C) or with exogenous TGF-β1 (B and D). (A) Cells cultured in control medium had an epithelial-like morphology in monolayer culture. (B) Morphological changes were observed after exposure to 2.0 ng/mL TGFβ1 for 72 h: multilayered aggregates formed in semiconfluent and confluent cultures and individual cells between aggregates became larger and developed filopodia and lamellipodia characteristic of a motile phenotype. (C and D) In control medium ZO1 was immunolocalized ( white speckled lines ) to lateral cell junctions ( arrowheads ). But immunostaining at cell borders was lost after culture with TGFβ1. Scale bar: 40 μm (A and B); 15 μm (C and D). Reproduced from Yang SP, Woolf AS, Yuan HT, Scott RJ, Risdon RA, O’Hare MJ, Winyard PJD. Potential biological role of transforming growth factor β1 in human congenital kidney malformations. Am J Pathol 2000;157:1633-1647.

Models of renal hypoplasia

The mildest type of kidney malformation is renal hypoplasia, where the kidney contains normally differentiated tissues but there are fewer nephrons than normal. Of note, even in overtly healthy, normotensive, humans there exists a wide range of nephron numbers, as assessed by counting glomeruli; the range is remarkable, from around half a million to two million [4]. Notably, individuals with a history of essential hypertension have, on average, half as many glomeruli per kidney [4]. Again, we can invoke genetic and environmental causes to explain variation in nephron number. For example, as shown in Fig. 1.11, glomerular numbers vary significantly between different strains of wild-type mice, and this may be attributed to yet-to-be defined genetic variation, e.g., C57BL6 has more glomeruli than FVB mice [5]. Moreover, within each strain, females tend to have more glomeruli than males [5], and one could hypothesize that testosterone inhibits nephrogenesis. In these studies, it was noted that the level of urinary albumin excretion in adult mice was inversely correlated with glomerular number [5], as shown in Fig. 1.12.

Fig. 1.11

Fig. 1.11 Glomerular counts in C57BL6 and FVB/N 18-week-old mice. (A) A reduction in glomerular number was observed in male (M) mice compared with strain-matched females (F); in addition, FVB/N had significantly lower number of glomeruli compared with sex-matched B6 mice. Similar changes were calculated when either (B) kidney or (C) body weight of the mice was taken into account (* P  < 0.05, ** P  < 0.01; *** P  < 0.001 between groups, n  = 6 in each group). All data are presented as means and standard error of the mean. Reproduced from Long DA, Kolatsi-Joannou M, Price KL, Dessapt-Baradez C, Papakrivopoulou E, Hubank M, Korstanje R, Gnudi L, Woolf AS. Albuminuria is associated with too few glomeruli and too much testosterone. Kidney Int 2013;83:1118–1129.

Fig. 1.12

Fig. 1.12 Albuminuria in C57BL6 and FVB/N mice. Overnight (A, C) albumin excretion and (B, D) albumin-to-creatinine ratios were evaluated in (A, B) 18-week-old and (C, D) 13-week-old adult mice. Data were log transformed before analysis and are presented as geometric means and confidence interval. There was a significant increase in urinary albumin in male (M) and female (F) FVB/N mice compared with sex-matched B6 animals. Within each strain, males had elevated albuminuria compared with females (** P  < 0.01, *** P  < 0.001 between groups). Reproduced from Long DA, Kolatsi-Joannou M, Price KL, Dessapt-Baradez C, Papakrivopoulou E, Hubank M, Korstanje R, Gnudi L, Woolf AS. Albuminuria is associated with too few glomeruli and too much testosterone. Kidney Int 2013;83:1118–1129.

A well-established example of an environmental perturbation able to generate renal hypoplasia is low protein diet initiated from the start of gestation in pregnant rats. Here, the offspring have significantly fewer glomeruli than normal. Moreover, this manipulation causes a markedly increased prevalence of apoptosis in the just-formed metanephros with a subsequent depletion in cell numbers [6]. Moreover, the low protein diet is associated with altered gene expression in the developing kidney [6]; the genes include those coding for PROX1, the ortholog of Drosophila transcription factor prospero, and cofilin-1, a regulator of the actin cytoskeleton. It has been reasoned that the effects of maternal low protein diet are in part mediated by downregulation of placental 11β-hydroxysteroid dehydrogenase-2 (HSD11B2), an enzyme degrading maternal corticosteroids [63]; this would lead to the embryo being exposed to high levels of glucocorticoids. Explants of mouse metanephric kidneys treated with dexamethasone, a glucocorticoid, show impaired overall organ growth as well as cystic change in the metanephric tubules [64]. This is associated with the upregulation of Indian hedgehog, a member of the SHH family, and addition of cyclopamine, a chemical that blocks HH signaling, ameliorates the malformation caused by dexamethasone [64]. An in vivo example of the plasticity in nephron numbers is provided by an experiment in which fetal sheep underwent uninephrectomy. Several weeks later, the remaining kidney had generated significantly more glomeruli than normal [65], perhaps in response to a yet-to-be defined circulating nephrogenic factor.

Urinary tract malformations

Probably, less of understood about the causes of urinary tract malformations, although in the last decade several genes have been found to be mutated in certain individuals with ureter or bladder anomalies. These genes code for transcription factors, smooth muscle contractile proteins, and neurogenic molecules, and the interested reader is directed elsewhere for a review of this topic [55]. As with the kidney, perhaps there are prospects for a degree of plasticity in urinary tract malformations. As an example, in a DNA (cytosine-5)-methyltransferase 1 mutant mouse model with a primary defect in bladder urothelial differentiation, a population of cells from the Wolffian duct migrate to the bladder and transdifferentiate to urothelium [66].

Summary

In this chapter, we have outlined the anatomy and molecular controls of kidney and urinary tract development. These processes are prone to error, with genetic and environmental factors playing their parts. We suggest that, by studying the pathobiology of renal tract malformations, clues will emerge to suggest pathways toward novel therapies which will restore the balance toward more normal development.

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Chapter 2: Renal organogenesis in the lymph node microenvironment

Maria Giovanna Francipane    McGowan Institute for Regenerative Medicine and Department of Pathology, University of Pittsburgh, Pittsburgh, PA, United States

Ri.MED Foundation, Palermo, Italy

Abstract

The shortage of organs for kidney transplantation has created an urgent need to find alternative treatment options. Recently, the combination of organoid cell culture technology with methodologies to fabricate three-dimensional scaffolds emerged as a promising approach to improve the function of several organs, including the kidney. An alternative approach is to urge organoids to mature into functional tissues by directly implanting them into a supportive niche in the host. Our unique approach takes advantage of the lymph node as a surrogate niche for functional kidney organogenesis. In the lymph node, primary mouse and human embryonic kidney fragments were nurtured and functionally matured. The lymph node also supported the engraftment and differentiation of organoid cultures derived from both mouse nephron progenitors and human-induced pluripotent stem cells that had been directed toward kidney fates. What makes the lymph node such a supportive site is its vascular-stromal network, which likely conveys specific cues to the developing transplanted tissue whether it be kidney or any other tissue. Further studies will be necessary to demonstrate the therapeutic efficacy of this approach.

In this chapter, I will briefly discuss current strategies and challenges for the engineering of a functional kidney, and discuss our strategy exploiting the lymph node microenvironment.

Keywords

Transplantation; Tissue engineering; Organoid; Kidney development; Lymph node; Vascularization

Acknowledgments

This work was supported by Ri.MED Foundation. We thank Lynda Guzik at the McGowan Institute Flow Cytometry Facility for proofreading the manuscript.

Funding

Supported by Ri.MED Foundation (M.G.F.).

Introduction

The kidney is a multifunctional organ which performs several essential physiological functions, including excretion of metabolic wastes, osmoregulation, and hormone secretion. Due to their complex function and intricate structure, renal tissues are markedly vulnerable to injury. It is known that the mammalian kidney is capable of post-injury growth, albeit in a limited capacity. It cannot make new nephrons, and although some regeneration events take place following acute kidney injury (AKI), they are exclusively confined to the tubular epithelium [1]. However, tubular regeneration is frequently incomplete and accompanied by fibrosis and long-term loss of function. Severe and repeated episodes of AKI lead to chronic kidney disease (CKD). End-stage renal failure, also known as an end-stage renal disease (ESRD), is the final, permanent stage of CKD. Two million people worldwide suffer from ESRD, and the number of patients diagnosed with this disease continues to increase at a rate of 5%–7% per year [2]. ESRD patients require dialysis or a kidney transplant to survive. While life-saving in the short-term, dialysis is not a long-term substitute for a human kidney, performing only about 10% of kidney function, and it is associated with increased left ventricular mass, intramyocardial cell fibrosis, and capillary loss [3, 4]. Orthotopic kidney transplantation is the only definitive treatment option for ESRD patients [5]. It is however a major surgical procedure that has risks both during and after the surgery. For example, immunosuppressive therapy, which is administered to almost all kidney transplant recipients to prevent acute rejection and the loss of the renal allograft, is associated with an increased incidence of cancer and viral opportunistic infections [6]. Graft rejection and post-transplant complications resulting from over-immunosuppression are not the only limitations for kidney transplantation. Shortage of kidney donors is still a major limitation [7]. Indeed, despite efforts to promote the practice of organ donation, the resources available in many countries are currently insufficient to meet medical needs. As such, there is a growing need to find better, alternative therapies to treat varying forms of kidney disease. One such avenue involves the field of regenerative medicine to either replace, restore, or regrow renal tissues.

Recently, tissue organoids have emerged as sophisticated models for studying development and disease, and as potential sources for creating organ substitutes [8], including the kidney. However, kidney engineering is a challenging task due to the complexity of an organ which contains at least 26 different types of cells organized in a unique structure. A series of challenges must be faced. First, the identification and selection of the best-suited stem/ progenitor cell type(s) for organoid generation. For years, the lack of definitive stem/progenitor cells in the adult kidney has been the major obstacle to renal repair and reconstruction [9]. With the development of the induced pluripotent stem cell (iPSC) technology, enabling the induction of pluripotency in mature somatic cells by treatment with defined factors [10], new opportunities to create patient-specific surrogate organs have emerged. Additional challenges of no less importance are the induction of substantial vascularization in kidney organoids in vitro, and the selection of a supportive transplantation site. This last objective is critical to allow the vascularized organoid to develop to functional maturity and perform important functions such as blood filtration, waste excretion, and essential substance reabsorption. We identified the lymph node—a secondary lymphoid organ which acts as a filter for foreign particles and cancer cells—as a surrogate niche for functional kidney organogenesis. The lymph node rapidly responds to accommodate increases in cellularity and has immediate vascular access, prerequisites for successful development of the kidney and its main function of blood filtration.

In this chapter, I will briefly discuss current strategies and challenges for engineering a functional kidney, and present our innovative strategy exploiting the lymph node. Our investigations seem to indicate that the lymph node might offer appropriate biophysical and biochemical cues to support kidney organogenesis.

Generation of tissue-engineered kidneys

Whole-organ tissue engineering has the potential to bridge the donor organ gap via the creation of artificial tissues by combining cells and matrices of natural or synthetic origin. Decellularization methodologies have been applied to produce whole organ-derived scaffolds by removing the cellular components while retaining all the necessary vascular and structural cues of the native organ [11]. However, the majority of described decellularization protocols show limitations in preserving the native structure of the extracellular matrix, mostly due to the harsh conditions required to remove cellular material [11]. At the moment, this technique only works with the heart in humans [12], and not the other major organs. Alternatively, supportive scaffolds can be produced from biomaterials including natural or synthetic polymers [13].

In the context of kidney, perhaps the most promising decellularization/recellularization strategy comes from the study of Song et al., which shows the generation of three-dimensional (3D) acellular renal scaffolds via perfusion decellularization of cadaveric rat, porcine, and human kidneys and their subsequent repopulation with endothelial and epithelial cells. Such kidney constructs were transplanted in an orthotopic position, and provided excretory function in vivo, following anastomosis to the recipient’s renal artery and vein [14]. Although promising, translation of this technology beyond proof of principle will first require the isolation, differentiation, and expansion of specific stem/progenitor cell types from clinically feasible sources.

A better understanding of the molecular basis of normal kidney development in the human, coupled with the breakthrough in research on pluripotent stem cells, has recently made the generation and derivation of patient-specific kidney cells a future possibility [15]. It is now well accepted that the mammalian kidney develops from the intermediate mesoderm through reciprocal interactions of the ureteric bud (UB) and the metanephric mesenchyme [16]. Three multipotent progenitor cell populations can be distinguished: 1) the Hoxb7   + UB progenitors, giving rise to the collecting duct and the ureter; 2) the Six2   + nephron progenitors, giving rise to most of the nephron; and 3) the FoxD1   + stromal progenitors, giving rise to multiple nonepithelial cell types within the kidney, including vascular progenitors, mural cells, and resident fibroblasts [15]. These progenitors are likely exhausted once nephrogenesis ceases—before birth in humans [17], and at postnatal day 3 in mice [18]. Tremendous effort has therefore been made to generate self-renewing populations of both nephron and UB progenitors from pluripotent stem cells for therapeutic purposes. Artificially created renal progenitors can generate 3D multicellular mini-organ structures, or organoids, in vitro, differentiating into kidney components following culture in appropriate media supplemented with recombinant growth factors. Markers of mature nephron components (podocytes, proximal tubules, loops of Henle, and distal tubules) are detected in nephron progenitor organoids [19–23]. In some cases, organoids also contain a renal interstitium and an endothelial network [19]. Similarly, under appropriate 3D culture conditions, UB progenitors can be expanded in vitro as branching UB organoids. Aggregating 3D-cultured nephron progenitors with UB organoids have recently been attempted in vitro, resulting in a reiterative process of branching morphogenesis and nephron induction, similar to kidney development [24, 25]. Despite these remarkable advancements, there is a current lack of full maturation, well-developed vascularization as well as a limitation in scalability and reproducibility [26]. Transcriptomic studies have often revealed the immaturity of kidney organoids in terms of differentiation and functional capabilities [19]. Obtaining a mature organoid is crucial to minimize the chances of unwanted tissue differentiation after transplantation and to guarantee the developing tissue is functional. Kidney function not only requires the coordinated action of various cell types organized into specific segments, but also necessitates a special 3D structure allowing the countercurrent mechanism to concentrate urine [27]. While a scaffold might help achieve a better spatial orientation of tubules, the self-organizing processes that build organoids cannot be easily matched by engineering approaches relying on a scaffold. Controlling self-organizing systems is inherently difficult because they follow intrinsic developmental programs, and small changes in the initial conditions inevitably lead to variable outcomes. In light of these limitations, scaffold-free organoid-based bioprinting emerged as a promising alternative [28].

Traditional transplantation sites for experimental kidney tissue engineering

The selection of the best cell type(s) for kidney organoid formation and the generation of a specific spatial and histo-topographic arrangement of nephron tubules are not the only challenges to overcome in kidney tissue engineering. Even assuming the most recent cell technologies would give rise to functional and well-aligned nephrons, where should they be transplanted? The possibility of implanting developing nephrons beneath the renal capsule of a postnatal or an adult host kidney has been studied since the early 90s [29]. However, this approach is not feasible in the clinic for at least two reasons. First, significant differences exist between murine and human kidney anatomy inasmuch as the human kidney capsule and parenchyma cannot be as easily separated as in rodents to permit cell transplantation [30]. Second, the fibrotic and inflammatory milieu of a chronically injured human kidney might not be an ideal niche for functional kidney organogenesis.

Organogenesis in an ectopic site was proposed as an alternative method to provide auxiliary organ function [31, 32]. Several ectopic sites were investigated in animals as transplantation sites for renal primordia (metanephroi), including the anterior eye chamber [33], the omentum [34], the retroperitoneal fat [35], the abdominal aorta [36], the paraaortic area [37], and the epididymis [38]. Metanephroi transplanted under the renal capsule [39] or in such ectopic sites [33–38] enlarged, become vascularized, formed glomeruli and tubules able to concentrate urine to a degree. However, the developing kidney would grow and survive for only a short time, unless an end-to-end anastomosis to the host ureters was performed [35, 36, 38]. Ureteroureterostomy between the transplant and native ureter prolonged short-term survival of anephric rats [35], and raised blood pressure in acutely hypotensive rats [38]. Although promising, these results point to several clinical limitations. Except for the omentum, none of the analyzed sites has the potential to reach the clinical practice. The greater omentum has already created significant interest for its potential as a site for pancreatic islet transplantation. A clinical trial investigating outcomes in patients who receive islet transplants to this site been launched [40]. However, major complications related to the use of the omentum include postoperative adhesions and intestinal obstruction [41, 42].

The lymph node microenvironment as an alternative site for kidney tissue engineering

The lymph node is a secondary lymphoid organ that has a unique ability to act as a bioreactor, allowing for substantial expansion of white blood cells during various conditions. This property is exploited by cancer cells, which often transit via the lymph node where they efficiently grow before disseminating to distant organs. Our group has used this growth-permissive environment to grow several tissues, including the kidney [43–51]. Following early kidney rudiment transplantation, robust kidney tissue forms in which nephron-like structures can be identified by molecular markers. We observed that host vasculature connects to the engrafted tissue allowing host blood flow through glomeruli, strongly suggesting the physiological function of the developing nephrons. Notable features of this engraftment technology make it a unique tool in which to study the organogenetic potential of pluripotent stem cell-derived kidney tissues and might have future clinical implications. In the paragraphs below, I will discuss our results of kidney organogenesis in this unique site.

Engraftment of mouse and human fetal primary tissues in the lymph node

By using fragments of green fluorescent protein (GFP) transgenic mouse metanephric (E14-15.5) kidneys, we first showed successful nephron maturation inside a host mouse lymph node [47, 48]. Renal tissues were harvested from C57BL/6 GFP   + transgenic embryos, isolated from UBs, minced, and injected directly into a single jejunal lymph node of adult wild-type C57BL/6 mice [47]. Following 3 weeks, recipient mice were sacrificed, lymph nodes were collected, and histologically examined [47]. Although some s-shaped bodies (glomerular and nephron precursors) could still be identified, the grafts contained several mature glomeruli, expressing CD31   + endothelial cells, and podoplanin   + podocytes [47]. Engrafting kidneys also showed cytokeratin-8 (K8) + rudimentary tubules as well as tubular erythropoietin expression [47].

By 6 weeks, mature nephrons consisting of mature glomeruli as well as renal tubules were observed [47]. Moreover, erythrocyte presence inside the mature capillary tuft indicated probable glomerular filtration ability [47].