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Novel functions of the macula densa in renal physiology and disease

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Novel functions of the macula densa in renal physiology and disease

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Content NOVEL FUNCTIONS OF THE MACULA DENSA IN RENAL PHYSIOLOGY AND
DISEASE

by

Urvi Nikhil Shroff, M.Sc. Biotechnology

A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(MEDICAL BIOLOGY)

August 2021

Copyright 2021 Urvi Nikhil Shroff
ii

Epigraph

“Superficially, it might be said that the function of the kidneys is to make urine; but in a more
considered view one can stay that the kidneys make the stuff of philosophy itself.”
Homer W. Smith, Lectures on the Kidney (1943)

iii

Dedication

To the quiet fortitude of women, their dogged persistence, and unwavering faith in each other.

To my family, of birth and of choice, to whom I owe a debt of gratitude.

iv

Acknowledgements

Dissertation Committee Members
Dr. Alicia McDonough (Chair)
Dr. János Peti-Peterdi (Mentor)
Dr. Laura Perin
PIBBS Medical Biology Program
Dr. Martin Kast (Program Director)
Domonique Walker (Student Advisor)

It takes a village for anyone to successfully complete their Ph.D. and my experience is no
different. First and foremost, I would like to express my immense gratitude to my Ph.D. Mentor,
Dr. János Peti-Peterdi for his guidance, help, and support throughout the last 5 years. I first had
the opportunity to work in his laboratory in Summer 2015 as a part of the Khorana Program for
Scholars and this experience played a formative role in my decision to pursue my doctoral studies
at the University of Southern California under his mentorship. Dr. Peti-Peterdi has been an
immense source of scientific, academic, and emotional support and encouragement throughout the
last several years. He provided unwavering support throughout my dissertation, encouraging me
to explore different aspects of my projects, and had faith in my abilities to successfully complete
the task at hand and I could not have asked for a more inspiring mentor than him. Dr. Peti-Peterdi
constantly encouraged me to apply for research fellowships, supported attending scientific
v

conferences and meetings while emphasizing maintaining a healthy work-life balance. His
immense patience as I learned various techniques and help with writing fellowship grants,
abstracts, and research articles are qualities that I will always appreciate. Dr. Peti-Peterdi’s
commitment to developing hypotheses-driven scientific projects continues to inspire me, and I will
take his teachings with me as I embark on the next stage of my professional and personal life.
Under his guidance, I have learned to be a better student and researcher, making this a truly
enriching experience.
I would like to express my deep gratitude to the members of my Dissertation Committee,
Dr. Alicia McDonough and Dr. Laura Perin. Over the last 4 years, they have provided valuable
insights and guidance, ensuring that I stayed on track to complete my dissertation. Their
constructive feedback, scientific input, and mentorship were instrumental in shaping my
hypotheses and research strategies. I would like to thank them for taking out time from their busy
schedules to guide me and for constantly pushing me to improve as a researcher. I would also like
to thank Dr. Sarah Hamm-Alvarez for providing vital ideas to better design my experiments.
My experience in the Peti-Peterdi laboratory has been particularly joyous due to all the
present and past laboratory members that I have had the pleasure to work with. Dr. Georgina
Gyarmati has been a wonderful colleague from whom I have learned a lot of scientific techniques
and skills, especially intravital imaging. Throughout my dissertation, Dr. Gyarmati helped me
troubleshoot experiments, discussed alternative strategies, and helped with data analysis. I am very
thankful for her support, friendship, and conversations, both scientific and otherwise. None of my
experiments would have been possible without the support of our laboratory manager, Audrey
Izuhara. She has been a source of comfort and mirth, always ready to help with any challenge that
I would face. In times of frustration or panic, when we ran out of reagents, or when I just needed
vi

to vent, Audrey was always there with a kind smile, a witty remark, and often, a bag of chips! I
would also like to thank Sachin Deepak for his help with my experiments and for maintaining the
mouse colonies for the past few months. I am also immensely grateful to our past laboratory
members, particularly Dr. Anne Riquier-Brison and Dr. Dorinne Desposito. Both Dr. Riquier-
Brison and Dr. Desposito helped me navigate the initial few years of my doctoral studies, teaching
me how to organize and plan experiments and I will always appreciate their generosity and
patience.
I would be remiss if I failed to mention the help, I have received from the PIBBS office-
Bami Andrada, Joyce Perez, Domonique Walker-, my entire PIBBS cohort, Dr. Ite Offringa, and
Dr. Martin Kast. I would also like to acknowledge my past mentors and professors- Mrs. Koshy
and Mrs. Bapat who instilled in me a love for biological sciences and Dr. Geetanjali Tomar, Dr.
Madhulika Dixit, Dr. Dominique Eladari, and Dr. Régine Chambrey for introducing and guiding
me as I began exploring a career in research.
I have been lucky and blessed to be surrounded by a wonderful group of friends as I worked
my way through graduate school. Dr. Krutika Deshpande, Dr. Rucha Bapat, and Dr. Kaivalya
Shevade - thank you for being wonderful sounding boards to bounce off ideas for experiments and
countless evenings filled with laughter. I could not have asked for better companions on this
journey through graduate school. To my best friend, Maitrayee Patil, thank you for understanding
me so completely, for your relentless support, for constantly pushing me outside my comfort zone,
and of course, for moving to Los Angeles and making it feel even more like home! Thank you also
to Shweta Pargaonkar, Tejas Narsule, Shagun Wazir, Samarth Syal, Sahil Joshi, Aakash Kabbin,
and Chaitanya Hulagabali for their friendship. Last but not the least, thank you to Sailesh Sidhwani
for always being there for me throughout the last 5 years. From helping me set up my first
vii

apartment in Los Angeles to being my travel partner on countless trips and adventures and patiently
listening as I narrated another day of failed experiments in the laboratory, Sailesh has been a
steadfast source of comfort and happiness.
I moved away from my home in India exactly 5 years ago to pursue my goal of earning a
Ph.D. and this would have been impossible without the unwavering support, help, and love I
received from my family - my parents Anita (and soon to be Dr. Anita!) and Nikhil and my siblings
Tanvi and Kaushik. No words can ever sufficiently describe how grateful and lucky I am to have
parents such as mine- they taught me the privilege and importance of receiving a well-rounded
education while giving me the freedom to make my own career choices. Despite being thousands
of miles away, my parents have been a constant source of inspiration, encouraging me during tough
times, supporting me as I tried to build a life in a country far from home, and making sure that I
was safe and healthy, and happy. Ma and Papa- thank you for your trust in me, for ensuring that I
never lacked for anything, for instilling in me the importance of hard work and perseverance, and
for being my North Star. My siblings have been a source of joy as I have watched them grow up
to be wonderful, accomplished (and very tall!) adults. Tanvi - thank you for amazing me with your
talents, for your warm spirit and maturity, and your passionate pursuit of a full life. Kaushik - your
dry wit, your clarity of thought, your quiet brilliance, and your sheer ability to persevere even
when the times get tough are qualities that I have always admired. Thank you both for making my
childhood full of laughter, for always having my back, and for holding down the fort when I moved
away.

-Urvi Nikhil Shroff
viii

Table of Contents
Epigraph .......................................................................................................................................... ii
Dedication ...................................................................................................................................... iii
Acknowledgements ........................................................................................................................ iv
List of Figures ................................................................................................................................ xi
Abbreviations ............................................................................................................................... xiii
Abstract ....................................................................................................................................... xvii
Chapter 1 ......................................................................................................................................... 1
Introduction ..................................................................................................................................... 1
1.1 The macula densa cell plaque ............................................................................................................. 1
1.2 Sensory functions of MD cells ............................................................................................................ 4
1.3 Regulation of renin release and glomerular filtration rate and renal hemodynamics via
tubuloglomerular feedback ....................................................................................................................... 5
1.4 The mammalian target of rapamycin signaling pathway .................................................................... 7
1.5 The Wnt/β-catenin signaling pathway ................................................................................................ 9
1.6 Crosstalk between the MAPK, mTOR, and Wnt/β-catenin signaling pathways .............................. 11
1.7 Genesis and scope of the current dissertation project ....................................................................... 12
1.8 References ......................................................................................................................................... 14
Chapter 2 ....................................................................................................................................... 19
A new view of macula densa cell protein synthesis
#
.................................................................... 19
2.1 Abstract ............................................................................................................................................. 19
2.2 Introduction ....................................................................................................................................... 20
2.3 Materials and Methods ...................................................................................................................... 22
2.3.1 Animals ...................................................................................................................................... 22
2.3.2 Treatments .................................................................................................................................. 23
2.3.3 Tissue processing and immunofluorescence .............................................................................. 23
2.3.4 Global protein synthesis assay using O-propargyl-puromycin fluorescence imaging and
quantification ...................................................................................................................................... 24
2.3.5 Intravital multiphoton microscopy ............................................................................................. 25
ix

2.3.6 Immunoblotting .......................................................................................................................... 26
2.3.7 Glomerular filtration rate measurements ................................................................................... 27
2.3.8 Blood pressure measurements .................................................................................................... 27
2.3.9 Statistical methods ..................................................................................................................... 27
2.4 Results ............................................................................................................................................... 28
2.4.1 Characterization of global protein synthesis in renal cell populations ....................................... 28
2.4.2 Generation and characterization of MD-mTOR
gof
mouse model ............................................... 31
2.4.3 Quantification of MD cell global protein synthesis in control and MD-mTOR
gof
mice ............. 34
2.4.4 Characterization of the autocrine effects of MD-mTOR signaling ............................................ 36
2.4.5 Blood pressure and kidney function in MD-mTOR
gof
mice ....................................................... 38
2.4.6 Intravital imaging of glomerular hemodynamics in MD-mTOR
gof
mice ................................... 38
2.4.7 Effects of MD mTOR signaling on renin expression ................................................................. 41
2.4.8 Changes in MD signaling in MD-mTOR
gof
mice ....................................................................... 42
2.5 Discussion ......................................................................................................................................... 45
2.6 References ......................................................................................................................................... 54
Chapter 3 ....................................................................................................................................... 58
Essential role of macula densa cell Wnt signaling in endogenous kidney tissue remodeling
*
..... 58
3.1 Abstract ............................................................................................................................................. 58
3.2 Introduction ....................................................................................................................................... 59
3.3 Materials and Methods ...................................................................................................................... 61
3.3.1 Animals ...................................................................................................................................... 61
3.3.2 Treatments .................................................................................................................................. 62
3.3.3 Tissue processing and immunofluorescence .............................................................................. 62
3.3.4 OPP labeling and quantification ................................................................................................ 63
3.3.5 Immunoblotting .......................................................................................................................... 63
3.3.6 Tissue clearing ........................................................................................................................... 64
3.3.7 Glomerular filtration rate measurement ..................................................................................... 65
3.3.8 Statistical methods ..................................................................................................................... 65
3.4 Results ............................................................................................................................................... 66
3.4.1 Characterization of Wnt/β-catenin signaling activity in macula densa cells ............................. 66
3.4.2 Generation and validation of MD-Wnt
lof
and MD-Wnt
gof
mouse models .................................. 68
3.4.3 Morphological and functional characterization of MD-Wnt
lof
and MD-Wnt
gof
mouse models .. 69
3.4.4 Paracrine effect of MD Wnt signaling ....................................................................................... 72
3.4.5 Regulation of MD cell protein synthesis by MD Wnt signaling ................................................ 74
x

3.4.6 Tissue remodeling activity of MD Wnt signaling ...................................................................... 76
3.5. Discussion ........................................................................................................................................ 78
3.6 References ......................................................................................................................................... 84
Chapter 4 ....................................................................................................................................... 87
Disease modifying role of macula densa cell Wnt signaling activity in diabetic kidney injury ... 87
4.1. Abstract ............................................................................................................................................ 87
4.2 Introduction ....................................................................................................................................... 88
4.3 Materials and Methods ...................................................................................................................... 90
4.3.1 Animals ...................................................................................................................................... 90
4.3.2 Treatments .................................................................................................................................. 91
4.3.3 Tissue processing and histology ................................................................................................ 91
4.3.4 Measurement of albuminuria and blood glucose ....................................................................... 92
4.4 Results ............................................................................................................................................... 92
4.4.1 Generation of diabetic kidney disease in control WT, MD-Wnt
lof
, and MD-Wnt
gof
mice ......... 92
4.4.2 Phenotypic and histological characterization of DKD mouse models ....................................... 93
4.4.3 Changes in albuminuria due to induction of DKD ..................................................................... 96
4.5 Discussion ......................................................................................................................................... 97
4.6 References ....................................................................................................................................... 101
Chapter 5 ..................................................................................................................................... 104
Conclusions and future directions ............................................................................................... 104
5.1 Summary of dissertation project ..................................................................................................... 104
5.2 Limitations and future directions .................................................................................................... 108
5.3 References ....................................................................................................................................... 110
Bibliography ............................................................................................................................... 112

xi

List of Figures

Figure 1.1. Structure of the nephron………………………………………………………………1
Figure 1.2. Structure of the renal corpuscle…………………………………………………….....2
Figure 1.3. Cell membrane ion transporter profile of MD cells……………………………………3
Figure 1.4. MD sensing and signaling pathways……………………………………………….....5
Figure 1.5. Schematic of the mTOR signaling pathway…………………………………………..8
Figure 1.6. Schematic of the Wnt/β-catenin signaling pathway………………………………… 10
Figure 1.7. Schematic depicting crosstalk between MAPK, mTOR, and Wnt pathways……… 11
Figure 2.1. Histological features of the Sox2-tdTomato mouse model………………………… 29
Figure 2.2. Quantitative visualization of protein synthesis activity in the kidney at the single-cell
level using O-propargyl puromycin-incorporation based fluorescence imaging………………...31
Figure 2.3. Validation of the expression of mTOR signaling elements in human and control and
MD-mTOR
gof
mouse kidney……………………………………………………………………..33
Figure 2.4. Quantification of MD cell global protein synthesis in control and MD-mTOR
gof
mice
using OPP-incorporation based fluorescence imaging…………………………………………...35
Figure 2.5. Autocrine and paracrine effects of upregulated MD mTOR signaling………………37
Figure 2.6. Intravital imaging of glomerular hemodynamics in MD-mTOR
gof
mice…………...40
Figure 2.7. Changes in renin expression in MD-mTOR
gof
mice………………………………….41
Figure 2.8. Changes in MD signaling in MD-mTOR
gof
mice……………………………………44
xii

Figure 3.1. Wnt/β-catenin signaling activity in macula densa cells……………………………..67
Figure 3.2. Validation of MD-Wnt
gof
and MD-Wnt
lof
mouse models……………………………69
Figure 3.3. Morphological and functional characterization of MD-Wnt
gof
and MD-Wnt
lof
mouse
models……………………………………………………………………………………………71
Figure 3.4. Paracrine effect of MD Wnt signaling on renin expression…………………………73
Figure 3.5. Quantification of MD cell protein synthesis in response to altered MD Wnt signaling
using O-propargyl-puromycin-incorporation based fluorescence imaging………………………75
Figure 3.6. Tissue remodeling effect of MD Wnt signaling……………………………………..77
Figure 4.1. Measurement of blood glucose levels in healthy and diabetic wildtype control, MD-
Wnt
lof
, and MD-Wnt
gof
mice……………………………………………………………………...93
Figure 4.2. Effects of genetic manipulations of MD Wnt signaling on kidney weight………….94
Figure 4.3. Histological characterization of kidney tissue in healthy and diabetic wildtype control,
MD-Wnt
lof
, and MD-Wnt
gof
mice………………………………………………………………...95
Figure 4.4. Measurement of albuminuria in response to altered MD Wnt signaling……………97
Figure 5.1. Summary of the current dissertation project………………………………………..104
Figure 5.2. Schematic depicting newly identified relevant MD cell signaling pathways………105
Figure 5.3. Schematic depicting novel role of MD cells in tissue remodeling…………………107

xiii

Abbreviations

4E-BP1- eIF4E binding protein 1
AA- afferent arteriole
ABC- active β-catenin
AC3- adenylate cyclase
ACR- albumin-to-creatinine ratio
AMDCC- Animal models of diabetes complications consortium
Ang II- angiotensin II
ATP- adenosine triphosphate
BABB- benzyl alcohol/benzyl benzoate
BP- blood pressure
BSA- bovine serum albumin
CCN1- CCN family member 1
CCN3- CCN family member 3
CHX- cycloheximide
CK1- casein kinase 1
CKD- chronic kidney disease
COX2- cyclooxygenase 2
CRD- cysteine-rich domain
CxCl14- C-X-C motif chemokine ligand 14
DC-destruction complex
Deptor- DEP domain containing mTOR interacting protein
DIC- differential interference contrast
DKD- diabetic kidney disease
DKK1- Dickkopf 1
DM- diabetes mellitus
DMSO- dimethyl sulfoxide
DN- diabetic nephropathy
DT- distal tubule
xiv

EA- efferent arteriole
eEF2- eukaryotic translation elongation factor 2
eGFP- enhanced green fluorescent protein
eIF3C- eukaryotic translation initiation factor 3C
eIF4E- eukaryotic translation initiation factor 4E
ER- endoplasmic reticulum
ERK1/2- extracellular-signal regulated kinase 1/2
ESRD- end-stage renal disease
FFPE- formalin-fixed paraffin-embedded
FZD- Frizzled
GBM- glomerular basement membrane
GC- glomerular capillary
GFP- green fluorescent protein
GFR-glomerular filtration rate
GLP 1- glucagon-like peptide 1
gof- gain-of-function
GPR91- G-protein coupled receptor 91
GSK3- glycogen synthase kinase 3
GSK3β- glycogen synthase kinase 3β
HPA- Human Protein Atlas
IC- intercalated cell
JG- juxtaglomerular
JGA- juxtaglomerular apparatus
LEF- lymphoid enhancer factor
lof- loss-of-function
LRP5/6- low-density lipoprotein receptor-related protein 5/6
LS- low salt
MAPK- mitogen activated protein kinase
MD- macula densa
Meis2- Meis homeobox 2
xv

mLST8- mammalian lethal with Sec13 protein 8
mPGES1- microsomal prostaglandin E2 synthase 1
MPM- multiphoton microscopy
mSIN1- mammalian stress-activated protein kinase interacting protein
mTmG- membrane targeted tdTomato membrane targeted eGFP
mTOR- mammalian target of rapamycin
mTORC1- mTOR complex 1
mTORC2- mTOR complex 2
NHE2- Na
+
/H
+
exchanger 2
NHE4- Na
+
/H
+
exchanger 4
NKCC2- Na
+
-K
+
-2Cl
-
cotransporter 2
nNOS- neuronal nitric oxide synthase
NO- nitric oxide
NS- normal salt
OPP- O-propargyl-puromycin
p70S6K- p70S6 kinase
p90S6K- p90S6 kinase
Pappa2- pappalysin 2
PAS- periodic acid Schiff
PBS- phosphate-buffered saline
PFA- paraformaldehyde
PGE2- prostaglandin E2
p-p70S6K- phospho-p70S6 kinase
PRAS40- proline-rich AKT substrate 40 kDa
Protor 1- protein observed with Rictor 1
PT- proximal tubule
PVDF- polyvinylidene difluoride
Rapa- rapamycin
Raptor- regulatory-associated protein of mTOR
RAS- renin-angiotensin system
xvi

RBC- red blood cell
RBCV- RBC velocity
RBF- renal blood flow
Rictor- rapamycin-insensitive companion of mTOR
RNAi- RNA interference
ROI- region of interest
ROMK- renal outer medullary K
+
channel
RPS6K- ribosomal protein subunit S6 kinase
SEM- standard error of mean
Sema3C- semaphorin 3C
SGLT1- sodium-glucose cotransporter 1
SGLT2- sodium-glucose cotransporter 2
Sox2- SRY-Box transcription factor 2
STZ- streptozotocin
TAL- thick ascending limb
TBST- tris-buffered saline with 0.1% Tween
TCF- T cell factor
tdTomato- tandem dimer Tomato
TGF- tubuloglomerular feedback
TSC2- tuberous sclerosis complex 2
VEGF-A- vascular endothelial growth factor-A
Veh- vehicle
WT- wildtype
WT1- Wilms’ tumor 1

xvii

Abstract

The kidney as a whole is composed of varied cellular populations, each with its specific
anatomy and function. Amongst these cell types, there exists a plaque of 20-25 specialized
epithelial cells, namely, the macula densa (MD) cells at the glomerular entrance of each nephron.
MD cells have long been recognized to play a critical role in regulating glomerular filtration rate
(GFR), renal blood flow (RBF), and renin release via tubuloglomerular feedback (TGF)
mechanisms, but the general understanding of the function of MD cells has been very limited. A
careful observation of the unique microanatomy of MD cells shows the abundance of endoplasmic
reticulum (ER), Golgi apparatus, and secretory granules in basal cell processes, which together
suggest that MD cells are endowed with an elaborate protein synthesis machinery towards the
basal surface facing the glomerulus. However, the complex three-dimensional architecture of the
glomerulus and the lack of a comprehensive MD cell research toolbox have made it difficult to
access and study the MD cells. The recent advancements in microscopy techniques, specifically
high resolution intravital multiphoton microscopy (MPM) and the development of MD-specific
transgenic mouse models now provide a new view of the mysterious MD cells and allow us to
carry out a detailed analysis of the unique features and functions of MD cells in the kidney.
The current dissertation project establishes and addresses the novel role of the high protein
synthesis and secretion activity of MD cells and its glomerulotrophic effect in mediating
endogenous tissue remodeling. Chapter 2 of the dissertation highlights the role of the mammalian
target of rapamycin (mTOR) signaling pathway in regulating MD cell protein synthesis activity as
well as its effect in regulating glomerular hemodynamics and renin expression. Chapters 3 and 4
xviii

address the effect of altered Wnt/β-catenin signaling activity in normal physiological as well as
pathophysiological settings.
In Chapter 2, we clearly demonstrate the importance of the mTOR pathway in upregulating
MD protein synthesis activity using a fluorescence imaging-based approach. Upregulated MD
mTOR signaling also significantly increased renin expression and classic MD signaling proteins,
resulting in robust increases in both GFR and RBF.
Chapter 3 pertains to the role of the MD Wnt signaling pathway in regulating glomerular
architecture and function. Using MD-specific mouse models to either upregulate or downregulate
Wnt signaling, we show several changes in the glomerular structure as well as GFR and renin
expression. Moreover, we also clearly demonstrate changes in specific cell populations-podocytes
and endothelial cells-which highlights the previously unknown role of MD cells in mediating
glomerular tissue remodeling.
Finally, in Chapter 4, we provide evidence for the disease modifying role of the MD Wnt
signaling pathway in the development and progression of diabetic nephropathy (DN).
In conclusion, the current dissertation provides a comprehensive overview of several newly
discovered aspects of MD cell function in the kidney and its role in health and disease including
in tissue regeneration and repair.

1

Chapter 1

Introduction

1.1 The macula densa cell plaque
The kidney is a highly complex organ, composed of over 20 different cell types that make
a concerted effort to serve the many kidney functions including the maintenance of body fluid and
electrolyte homeostasis and the regulation of blood pressure (BP) (1). The functional unit of the
kidney is the nephron; the human kidney has, on average, 1 million nephrons. Each nephron begins
with the renal corpuscle which then leads into a series of tubular segments (Fig. 1.1) (2).
Figure 1.1. Structure of the nephron
(2)
.
The renal corpuscle consists of the glomerulus and the Bowman’s capsule- the glomerular
structure consists of an afferent arteriole (AA) that leads into a tuft of glomerular capillaries (GCs)
and lead out as the efferent arteriole (EA) while the Bowman’s capsule surrounds the glomerulus.
2

The major tubular segments begin with the proximal tubule (PT), followed by the thin descending
limb and thick ascending limb (TAL) of the loop of Henle, and finally the distal tubule (DT) and
collecting duct system. Each tubular segment is composed of a single layer of highly specialized,
polarized epithelial cells that mediate the specific function of that tubular segment.
The structure of the nephron is such that the TAL of each nephron loops back towards its
parent glomerulus. At the end of the TAL where the nephron loops back towards the glomerulus,
there exist a plaque of approximately 20-25 specialized cells known as the “macula densa (MD)
cells” strategically positioned at the glomerular vascular pole (Figs. 1.1 and 1.2). MD cells form
the tubular component of the juxtaglomerular apparatus (JGA) along with the juxtaglomerular (JG)
cells of the AA and the extraglomerular mesangial cells (3).
Figure 1.2. Structure of the renal corpuscle
(4)
.
First identified by Golgi more than a century ago (5), the JGA is a hemodynamic filtration
unit that is a crucial site for the regulation of body salt and fluid balance and BP (6, 7). Within the
3

JGA, the MD cell plaque was first described by Zimmerman in 1930 based on the tight packing of
nuclei of tubular cells as compared to other DT cells (8). Early studies demonstrated several
distinctive features of these cells- closely packed, tall cells with a large apical nucleus, several
short ovoid mitochondria, extensive Golgi apparatus, and cytoplasmic processes at the basilar
surface (7). Functionally, MD cells act as chief sensors of the tubular filtrate salt concentration,
flow rate, and the local tissue environment. The apical surface of MD cells faces the tubular lumen
and is exposed to differing levels of several ions and metabolites present in the filtrate. On the
other hand, the basal surface of MD cells faces the extraglomerular mesangium, the JG cells, and
the glomerulus (3).
Cell membrane transporter profiling of MD cells showed that on the apical surface MD
cells possess a Na
+
:K
+
:2Cl
-
cotransporter 2 (NKCC2) (9, 10), a Na
+
/H
+
exchanger 2 (NHE2) (11),
a H
+
/K
+
-ATPase (12), and a renal outer medullary K
+
(ROMK) channel (13) while the basal
membrane expresses a Na
+
/H
+
exchanger 4 (NHE4) (11), a 380 pS maxi-anion channel (14) and
Figure 1.3. Cell membrane ion transporter profile of MD cells
(17)
.
4

Cl
-
and Ca
2+
channels (15). The polarized NHE2/NHE4 arrangement (11), the extremely low
activity of the basolateral Na
+
/K
+
-ATPase (16), and the alternative usage of the apical H
+
/K
+
-
ATPase (12) are unique features of MD cells that serve their sensory functions (17).

1.2 Sensory functions of MD cells
The primary sensory function of MD cells is to detect alterations in tubular fluid [NaCl]
and flow rate via its apical transporters- NKCC2 and NHE2, with NKCC2 being the primary source
of NaCl entry into the cell (10, 11, 17). MD cells express the high-affinity B isoform of NKCC2
which operates in the range of 20-60 mM [NaCl] (9, 18). NKCC2 activity requires continuous
availability of K
+
which is met by the apical activity of the ROMK channel (19). The apical NHE2
transporter operates in the range of 20-150 mM [NaCl] and is responsible for the alkalization of
the MD cells in response to increasing luminal [NaCl] (17). An increasing luminal NaCl
concentration also results in the depolarization of MD cells most likely due to the egress of Cl
-
from the basolateral Cl
-
channel (15). The basolateral NHE4 is activated in response to cell
shrinkage due to an increase in filtrate osmolality, suggesting a role for NHE4 in regulating cell
volume (11). Finally, as mentioned above, MD cells are unique in that they have an extremely low
level of the otherwise ubiquitous basolateral Na
+
/K
+
-ATPase and instead express an apical H
+
/K
+
-
ATPase to maintain an electrochemical gradient across the membrane (6, 12, 16, 17). With respect
to the detection of tubular fluid flow rate, it has been demonstrated that in response to elevated
fluid flow rate, cytosolic Ca
2+
increases significantly and this flow-sensing ability of MD cells is
attributable to the presence of a prominent apical primary cilium (20).
5

In addition to being an important sensor of tubular [NaCl], recent studies from several
laboratories including ours have shown that MD cells can also sense specific local environmental
factors including metabolites (21). For example, MD cells can sense the Krebs cycle intermediate
succinate using the apical G-protein coupled receptor 91 (GPR91) (22) and express olfactory G
protein and the olfactory isoform of adenylate cyclase 3 (AC3) (23).

1.3 Regulation of renin release and glomerular filtration rate and renal hemodynamics via
tubuloglomerular feedback
MD cells play a critical role in regulating renal blood flow (RBF) and glomerular filtration
rate (GFR) via the tubuloglomerular feedback (TGF) as well as renin release from the JG cells (6).
The composition of the tubular filtrate is altered by the action of various transporters along the
different tubular segments, with the bulk of the reabsorption occurring in the PT (3). Several
Figure 1.4. MD sensing and signaling pathways
(6)

6

factors determine the [NaCl] in the filtrate as it reaches the MD cells including renal perfusion
pressure, tubular fluid flow rate, [NaCl] reabsorption in PT segments, neurotransmitters of the
sympathetic nervous system, levels of locally acting hormones, etc. (6). A decrease in filtrate
[NaCl] detected by MD cells results in the release of renin from the JG cells of the AA via the
activation of a series of intracellular signaling pathways. Low tubular [NaCl] promotes the
phosphorylation and activation of 2 important mitogen activated protein kinases (MAPKs)-
extracellular-signal regulated kinase 1/2 (ERK1/2) and p38 along with 2 enzymes-cyclooxygenase
2 (COX2) (24) and microsomal prostaglandin E2 synthase 1 (mPGES1)- that results in the
production of the autocoid prostaglandin E2 (PGE2) (25).
The synthesized PGE2 is released from the MD basal surface and it acts on the EP2 and
EP4 receptors of the JG cells, triggering the release of renin from storage granules (6, 26). In cases
of elevated filtrate [NaCl], MD cells are known to release ATP from their basolateral membrane
380 pS maxi-anion channel which binds to purinergic receptors present on JG cells and inhibits
renin release due to the elevation of intracellular Ca
2+
(6, 14, 27, 28) (Fig. 1.4).
In addition to regulating renin release, MD cells are also involved in regulating renal and
glomerular hemodynamics via the TGF mechanism. The TGF system senses changes in the
delivery of filtrate to the DT and initiates a cascade of reactions to ensure autoregulation of GFR
and RBF. An increase in the tubular filtrate [NaCl] to the DT detected by the MD increases
preglomerular resistance, thereby reducing RBF and GFR and normalizing [NaCl] delivery to the
MD. The apical NKCC2 detects increases in tubular [NaCl], ultimately resulting in the basolateral
release of ATP and adenosine which trigger elevations of intracellular Ca
2+
of the AA and
mesangial cells, resulting in vasoconstriction of the AA and the glomerular tuft (17). The
combination of the TGF and the myogenic mechanism (alterations in vascular smooth muscle tone
7

of the AA) ensures constant delivery of tubular filtrate to the distal nephron, thereby maintaining
homeostasis (3).

1.4 The mammalian target of rapamycin signaling pathway
The specialized microanatomy of MD cells with the basal localization of various secretory
and protein synthetic organelles suggests that MD cells are endowed with a robust protein
synthesis machinery (7). The mammalian target of rapamycin (mTOR) signaling pathway is a
central regulator of cell metabolism, proliferation, and growth. The mTOR pathway is responsive
to both extracellular signals like growth factors, glucose, and amino acids and intracellular signals
like cellular energy and oxygen status (30). The mTOR protein, a 289 kDa serine-threonine kinase,
forms the core of 2 protein complexes- mTOR complex 1 (mTORC1) and mTOR complex 2
(mTORC2). The mTORC1 complex consists of 5 proteins- mTOR, regulatory-associated protein
of mTOR (Raptor) (31), mammalian lethal with Sec13 protein 8 (mLST8) (30), DEP domain
containing mTOR interacting protein (Deptor) (32), and proline-rich AKT substrate 40 kDa
(PRAS40) (33) and while the mTORC2 complex consists of 6 proteins- mTOR, rapamycin-
insensitive companion of mTOR (Rictor), mammalian stress-activated protein kinase interacting
protein (mSIN1) (35), protein observed with Rictor 1 (Protor 1), mLST8 and Deptor (30) (Fig.
1.5).

8

Figure 1.5. Schematic of the mTOR signaling pathway
(34)
.

mTORC1 is a positive regulator of several anabolic processes- protein synthesis, lipid
synthesis, and mitochondrial biogenesis while acting as a potent inhibitor of catabolic processes
including autophagy (30). With respect to protein synthesis, the mTORC1 complex is a positive
regulator of protein synthesis, mRNA translation, and ribosome biogenesis. The mTORC1
complex phosphorylates and activates the p70S6 kinase (p70S6K) ribosomal protein while
inhibiting the eukaryotic translation initiation factor 4E (eIF4E)-binding protein 1 (4E-BP1) (30).
Activation of p70S6K increases mRNA biogenesis and cap-dependent elongation while the
inhibition of the 4E-BP1 results in the promotion of cap-dependent translation by eIF4E (36, 37).
Finally, the mTORC1 complex also positively regulates ribosome biogenesis by promoting the
transcription of ribosomal RNA (38-40).
9

1.5 The Wnt/β-catenin signaling pathway
Preliminary data from our laboratory has shown that MD cells have a high level of Wnt/β-
catenin signaling activity (unpublished data). The canonical Wnt/β-catenin signaling pathway is
primarily involved in developmental and homeostatic processes via the action of a family of
growth-promoting factors-the Wnt proteins (41). Wnt proteins are lipid-modified cysteine-rich 40
kDa proteins that mediate their effects over a short range by binding to a receptor complex
consisting of Frizzled (FZD) and low-density lipoprotein receptor-related protein 5/6 (LRP5/6)
(42, 43). The lipid modification on Wnt proteins plays a key role in binding to the cysteine-rich
domain (CRD) of the FZD co-receptor (44). The binding of the Wnt ligand to its cognate receptor
complex triggers the cytoplasmic stabilization of β-catenin which is a part of the destruction
complex (DC). When the receptor complex is in the unbound state, the β-catenin in the DC is
phosphorylated by kinases in the DC- casein kinase 1 (CK1) and glycogen synthase kinase 3β
(GSK3β). The phosphorylated β-catenin undergoes ubiquitination and degradation by the
proteosome. Conversely, when the receptor complex is engaged, the ubiquitination of β-catenin is
inhibited, saturating the DC with β-catenin. The resulting accumulation of newly synthesized β-
catenin is free to translocate to the nucleus where it mediates the transcription of several target
10

genes along with the T cell factor (TCF)/lymphoid enhancer factor (LEF) family of transcription
factors (40) (Fig. 1.6).
Figure 1.6. Schematic of the Wnt/β-catenin signaling pathway
(41)
.
11

1.6 Crosstalk between the MAPK, mTOR, and Wnt/β-catenin signaling pathways
As mentioned previously, the MAPK pathway plays an important role in mediating the
function of MD cells in the kidney (6). Additionally, as discussed in this thesis, the mTOR and
Wnt signaling pathways also display a high level of activity within the MD cells. Several points
of integration exist between the MAPK, mTOR, and Wnt signaling pathways (Fig. 1.7).
Figure 1.7. Schematic depicting crosstalk between MAPK, mTOR, and Wnt pathways.
Created with Biorender.com
12

Activated ERK1/2 is known to phosphorylate and inhibit tuberous sclerosis complex 2
(TSC2), thereby resulting in increased mTORC1 activity (45). Additionally, ERK1/2-induced
phosphorylation of Raptor can also promote mTORC1-induced phosphorylation of 4E-BP1 (46).
ERK1/2 can also independently activate p90S6 kinase (p90S6K) while the role of mTORC1 in the
activation of p70S6K has been well-established (47, 48).
The Wnt/β-catenin pathway is known to interact with the mTOR signaling pathway via its
effect on GSK3β. Activation of Wnt signaling inhibits GSK3β-mediated phosphorylation of TSC2.
This bifurcation at the level of GSK3β allows the Wnt signaling pathway to regulate gene
transcription via β-catenin and protein translation via the mTOR pathway (49). Thus, each of these
signaling pathways is integrated at specific points and can play an important role in regulating
signaling activity (50).

1.7 Genesis and scope of the current dissertation project
Preliminary evidence from our laboratory suggests an important role for the mTOR and
Wnt/β-catenin signaling pathways in regulating MD cell function (unpublished data).
Immunohistochemistry data from the Human Protein Atlas (HPA) database (51) shows the high
level and cell-specific expression of several mTOR signaling proteins including TSC2 and Deptor
as well as downstream targets like p70S6K, eukaryotic translation initiation factor 3C (eIF3C),
and eukaryotic translation elongation factor 2 (eEF2) in the human MD. Additionally,
immunofluorescence labeling for TSC2 on mouse kidney tissue similarly reflected a high-level
expression in the MD cell plaque, underscoring the importance of this pathway in MD cells. Using
the WntGFP reporter mouse model (in which intensity of green fluorescent protein (GFP)
13

expression is directly proportional to Wnt signaling activity) (52) we observed that MD cells have
the highest Wnt signaling activity based on the intense GFP expression within MD cells. This was
further corroborated via immunofluorescence labeling for active-β-catenin (ABC) (the final
effector of this pathway) (42) on mouse kidney tissue and MD cell gene profile analysis via
RNASeq (unpublished data) which showed high expression of Wnt10a and numerous other Wnt
target genes. Based on these data, I aimed to determine the role of these specific signaling pathways
in the regulation of MD cell function and its paracrine effects on renal and glomerular structure
and function.
The central hypothesis of the current dissertation project is that as part of their novel
functions, MD cells are chief organizers of glomerular remodeling and control local mesenchymal
and endothelial progenitor cells via the synthesis and secretion of angiogenic and tissue remodeling
peptides. Moreover, MD cells feature a high level of protein synthesis for their secretory function
that is regulated by the mTOR and Wnt/β-catenin signaling pathways. To test this central
hypothesis, the current dissertation project was divided into three Specific Aims (SA):
SA1. To elucidate the key autocrine role of MD mTOR and Wnt/β-catenin signaling
pathways in regulating MD cell protein synthesis.
As a part of SA1, I aimed to elucidate the role of the MD mTOR and Wnt/β-catenin
signaling pathways in regulating the MD protein synthesis and its effect on glomerular architecture
and function. Using novel MD-specific genetic mouse models and state-of-the-art confocal and
multiphoton microscopy (MPM) techniques, I aimed to quantify changes in MD protein synthesis
activity in response to alterations in the activity of these two signaling cascades. The results from
these experiments are the focus of Chapters 2 and 3.
14

SA2. To test the role of MD mTOR and Wnt/β-catenin signaling pathways in the paracrine
control of glomerular structure and physiological function.
SA2 is targeted towards understanding both, the autocrine and paracrine effect of the MD
Wnt/β-catenin pathway on glomerular structure and function. Three-dimensional tissue clearing
techniques were used to study changes in the density of specific renal cell populations while
changes in renal function were quantified by measuring GFR and RBF. The results from these
experiments have been summarized in Chapters 2 and 3.
SA3. To provide evidence for the therapeutic relevance of MD Wnt/β-catenin signaling in a
glomerular disease model.
Finally, in SA3, I explored the role of the MD Wnt/ β-catenin signaling pathway in the
development and progression of renal pathophysiology. I used a well-established mouse model of
diabetic nephropathy (DN) to determine the role of the MD Wnt signaling pathway in the
pathophysiological context, as described in Chapter 4.
Taken together, these aims encompass the overarching goal of better understanding the
novel roles of MD cells in kidney tissue remodeling and disease.

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#
This chapter is modified from my first author original research article “Shroff UN, Gyarmati G, Izuhara A, Deepak S, and Peti-
Peterdi J. A new view of macula densa cell protein synthesis (submitted to AJP Renal F-00222-2021)”. The methods appear in my
first author book chapter “Shroff UN, Schiessl IM, Gyarmati G, Riquier-Brison A, Peti-Peterdi J. Novel fluorescence techniques
to quantitate renal cell biology. Methods Cell Biol 154: 85-107, 2019”.
19

Chapter 2

A new view of macula densa cell protein synthesis
#

2.1 Abstract

Macula densa (MD) cells, a chief sensory cell type in the nephron, are endowed with unique
microanatomical features including a high density of protein synthetic organelles and secretory
vesicles in basal cell processes (“maculapodia”) that suggest a so far unknown high rate of MD
protein synthesis. The present study aimed to explore the rate and regulation of MD protein
synthesis and their effects on glomerular function using novel transgenic mouse models, newly
established fluorescence cell biology techniques, and intravital microscopy. SRY-Box
transcription factor 2 (Sox2)-tdTomato kidney tissue sections and the O-propargyl puromycin
(OPP)-incorporation based fluorescence imaging assay showed that MD cells have the highest
level of protein synthesis within the kidney cortex followed by intercalated cells (ICs) and
podocytes. Genetic gain-of-function (gof) of mammalian target of rapamycin (mTOR) signaling
specifically in MD cells (in MD-mTOR
gof
mice) or their physiological activation by low salt (LS)
diet resulted in further significant increases in the synthesis of MD proteins. Specifically, these
included both classic and recently identified MD-specific proteins such as cyclooxygenase 2
(COX2), microsomal prostaglandin E2 synthase 1 (mPGES1), and pappalysin 2 (Pappa2),
respectively along with renin. Intravital imaging of the kidney using multiphoton microscopy
20

(MPM) showed significant increases in afferent (AA) and efferent arteriole (EA) and glomerular
capillary (GC) diameters and blood flow in MD-mTOR
gof
mice coupled with elevated glomerular
filtration rate (GFR). The presently identified high rate of MD protein synthesis that is regulated
by mTOR signaling is a novel component of the physiological activation and glomerular
hemodynamic regulatory functions of MD cells that remains to be fully characterized.
Keywords: macula densa, protein synthesis, glomerular filtration rate, renin
2.2 Introduction
Macula densa (MD) cells, the tubular component of the juxtaglomerular apparatus (JGA),
are crucial for the regulation of renal and glomerular hemodynamics via the tubuloglomerular
feedback (TGF) loop as well as renin release from the neighboring juxtaglomerular (JG) cells (1).
Strategically positioned at the glomerular vascular pole, the MD cell plaque consists of
approximately 20-25 highly specialized epithelial cells, present at the end of the thick ascending
limb (TAL) of the nephron (2). MD cells have a distinctive polarized morphology- the apical
surface includes a prominent primary cilium (3, 4) and a large nucleus (2) while the basolateral
surface is densely packed with secretory organelles (2) and endowed with a newly identified cell
processes network called “maculapodia” (5). These unique microanatomical features play a critical
role in mediating the traditional function of MD cells as sensors of tubular salt, flow (6), and other
local tissue environmental factors including metabolites (7-9). A decrease in tubular salt
concentration, detected by the apical Na
+
-K
+
-2Cl
-
cotransporter 2 (NKCC2), initiates an
intracellular signaling cascade of mitogen activated protein kinases (MAPKs)- p38 and
extracellular-signal regulated kinase 1/2 (ERK1/2) (1, 6, 10)- along with the enzymes-
cyclooxygenase 2 (COX2) (10) and microsomal prostaglandin E2 synthase 1 (mPGES1) (11, 12),
21

resulting in the release of the autocoid prostaglandin E2 (PGE2) which in turn triggers renin release
from the JG cells (13). The end result of the activation of this signaling cascade (the renin-
angiotensin system (RAS)) is the increase in the production of angiotensin II (Ang II). Ang II in
turn acts on many organ systems to increase extracellular fluid volume and blood pressure (BP),
and restore body fluid and electrolyte balance. These Ang II actions include in the kidney the
vasoconstriction of the AA, increased reabsorption of NaCl from the tubular filtrate as well as
promoting aldosterone activity on the distal tubule (DT)-collecting duct to further facilitate tubular
fluid NaCl reabsorption and to inhibit natriuresis (2).
The various renal cell populations synthesize cell-specific structural proteins, peptides, and
hormones to maintain their structural integrity and mediate their functions. For example, podocytes
are potent sources of vascular endothelial growth factor-A (VEGF-A) (14, 15), interstitial
fibroblasts produce erythropoietin (16), and JG cells secrete renin (17). Cellular protein synthesis
is tightly regulated at several checkpoints- transcription, translation, and degradation- and involves
various organelles including the endoplasmic reticulum (ER), Golgi apparatus, and the
mitochondria in order to maintain proteostasis (18, 19). A central regulator of cellular protein
synthesis and metabolism, the mammalian target of rapamycin (mTOR) signaling cascade is
activated by amino acids and growth factors, while it is inhibited during oxygen and nutrient
deficiency. Activated mTOR complex 1 (mTORC1) acts on several downstream targets including
the ribosomal kinase p70S6 kinase (p70S6K), mRNA translation proteins like eukaryotic
translation initiation factor 4E (eIF4E), and eukaryotic translation elongation factor 2 (eEF2),
thereby resulting in increased mRNA translation, ribosome biogenesis, and protein synthesis. In
addition to protein synthesis and cellular bioenergetics, the mTOR pathway also plays a crucial
role in regulating cellular proliferation and hypertrophy (20, 21). Regulated protein synthesis is
22

fundamental to the proper functioning of a specific cell type. Dysregulation of ER function and
protein biosynthesis due to stress factors, hypoxia, accumulation of misfolded proteins, and other
pathogenic insults are known to be involved in the pathogenesis of several diseases including
diabetic nephropathy (DN), nephrotic syndrome, cardiorenal syndrome, and Alzheimer’s disease
(22-24).
While the morphological features of MD cells suggest the presence of a robust protein
synthesis machinery at the basal surface (2), the relative inaccessibility of MD cells and complex
architecture of the JGA has limited our understanding of the protein synthesis and secretion
capacity of MD cells using conventional histology techniques. The recent development of novel
fluorescence techniques including an imaging-based, single-cell level global protein synthesis
assay and transgenic mouse models (25-28) has made it possible to extensively characterize and
quantify specific aspects of renal cell biology.
The present study aimed to characterize the rate and regulatory mechanisms of MD protein
synthesis using newly developed fluorescence imaging tools and MD-specific genetic mouse
models. In addition, our study also addressed the autocrine and paracrine effects of enhanced MD
cell protein synthesis on renin expression and glomerular hemodynamics.

2.3 Materials and Methods
2.3.1 Animals
All animal protocols were approved by the Institutional Animal Care and Use Committee
at the University of Southern California. Tamoxifen-inducible, conditional macula densa (MD)
cell-specific mammalian target of rapamycin (mTOR) gain-of-function (gof) (MD-mTOR
gof
) mice
23

on a C57BL6/J background were generated by intercrossing nNOS/CreERT2 and TSC2/fl mice
(exons 2-4 of Tsc2 are flanked by loxP sites) (Jackson Laboratory, Bar Harbor, ME) (29). MD-
mTOR
gof
mice were further backcrossed with the two-color fluorescent mTmG/fl reporter mice
(Jackson Laboratory) for the expression of the cell membrane targeted tandem dimer Tomato (mT,
tdTomato) and enhanced green fluorescent protein (mG, eGFP) such that MD cells specifically
express eGFP whereas all other cells express tdTomato (30). Tamoxifen-inducible, conditional
MD-GFP mice on a C57BL6/J background were generated by intercrossing nNOS/CreERT2 and
mTmG/fl reporter mice (Jackson Laboratory) as described before (5, 30). Constitutive Sox2-
tdTomato mice on a C57BL6/J background were generated by intercrossing SRY-Box
transcription factor 2 (Sox2)/Cre (31) and tdTomato/fl mice (Jackson Laboratory) for ubiquitous
expression of tdTomato in all renal cell types.
2.3.2 Treatments
Tamoxifen administration (75 mg/kg body weight, Alfa Aesar, Harverhill, MA) was via
oral gavage for a total of 3 times every alternate day. Low salt (LS) diet (TD 90228, Harlan Teklad,
Madison, WI) treatment was carried out for 2 weeks ad libitum for a subset of mice. For rapamycin
(Rapa) treatment, mice were injected intraperitoneally with Rapa (8 mg/kg body weight, Alfa
Aesar) every alternate day for 2 weeks by dissolving Rapa in ethanol followed by dilution in filter-
sterilized vehicle (0.9% NaCl, 5% PEG and 5% Tween-80) as described previously (32).
Cycloheximide (CHX) (Sigma Aldrich, St. Louis, MO) treatment was carried out by injecting mice
intraperitoneally with 300 µl of 16 mg/ml CHX as described before (33).
2.3.3 Tissue processing and immunofluorescence
Mice were anaesthetized using a combination of ketamine (100 mg/kg body weight) and
xylazine (10 mg/kg body weight) followed by perfusion with ice-cold 1X phosphate-buffered
24

saline (PBS) and 4% paraformaldehyde (PFA) for 2 minutes using a peristaltic pump. Tissue was
harvested and fixed in 4% PFA for 2 hours at room temperature. For frozen tissue blocks, tissue
was kept overnight in 30% sucrose at 4ºC for cryoprotection and then embedded in Tissue-Tek
O.C.T compound (Sakura Finetek, Torrance, CA) and flash frozen. Cryosections were cut at 25
µm, washed with 1X PBS, and mounted using DAPI-containing VectaShield mounting media
(Vector Laboratories Inc., Burlingame, CA). For paraffin tissue blocks, tissue was dehydrated and
embedded in paraffin wax and then sectioned 8 µm thick. For immunofluorescence, tissue sections
were treated with 0.5% Triton for 15 minutes followed by heat-induced antigen retrieval by boiling
sections in sodium citrate buffer (pH 6.0) for 10 minutes. To reduce non-specific binding of
antibodies, sections were blocked for 30 minutes with normal donkey serum (1:20). Primary and
secondary antibodies were applied sequentially overnight at 4ºC and for 1 hour at room
temperature, respectively, and mounted using DAPI-containing VectaShield mounting
media. Primary antibodies and dilutions were as follows: neuronal nitric oxide synthase (nNOS)
(Santa Cruz Biotechnology, sc-648, 1:100), renin (Anaspec Inc., AS-54371, 1:100), H-ATPase
(generous gift from Dr. Mark Knepper (34), 1:50), p57 (Santa Cruz Biotechnology, sc-8298,
1:100) and tuberous sclerosis complex 2 (TSC2) (Cell Signaling Technology, CST3612, 1:50).
AlexaFluor 594 and 647-conjugated secondary antibodies (Thermo Fisher Scientific, Waltham,
MA) were all used at 1:500 dilution. Sections were examined with Leica TCS SP8 DIVE (Leica
Microsystems, Wetzlar, Germany) confocal/multiphoton laser scanning microscope system as
described previously (5).
2.3.4 Global protein synthesis assay using O-propargyl-puromycin fluorescence imaging
and quantification
For O-propargyl-puromycin (OPP) labeling (Thermo Fisher Scientific), mice were injected
intraperitoneally with 25 µl of 20 mM OPP dissolved in dimethyl sulfoxide (DMSO) 1 hour before
25

tissue harvest and staining as described previously (25, 35). For OPP labeling, sections were
developed using the AlexaFluor 594 Click-iT OPP Protein Synthesis kit (Thermo Fisher Scientific)
according to the manufacturer’s instructions. OPP fluorescence intensity was quantified by
imaging all tissue sections using identical laser power and confocal microscopy settings.
Fluorescence intensity was measured by placing 10 regions of interest (ROIs) across the entire
MD plaque (FOPP) and normalized to the fluorescence intensity of red blood cells (RBCs) (FRBC).
The average FOPP of 5-10 different representative MD cell plaques was quantified in each tissue
section.
2.3.5 Intravital multiphoton microscopy
For intravital imaging of control MD-GFP and MD-mTOR
gof
mice, animals were
anesthetized with 1-4% isoflurane and the SomnoSuite low-flow anesthesia system (Kent
Scientific, Torrington, CT), and the left kidney was exteriorized via a small incision. Mice were
injected with Alexa Fluor 680-conjugated bovine serum albumin (BSA) (Thermo Fisher Scientific)
retro-orbitally to label the circulating plasma and placed on the stage of an inverted microscope,
with the exteriorized kidney bathed in a coverslip-bottomed chamber containing normal saline as
described previously (25). Body temperature was maintained with a homeothermic blanket system
(Harvard Apparatus, Holliston, MA) throughout the imaging session. Intravital imaging was
performed using a Leica SP8 DIVE multiphoton microscope (Leica Microsystems) with a 40X
Leica water-immersion objective (numerical aperture 1.1) and a Chameleon Discovery laser
system (Coherent Inc., Santa Clara, CA) at 960 nm and external Leica 4Tune spectral hybrid
detectors (emission at 530  20 nm for mG and 590  20 nm for mT). Changes in glomerular
hemodynamics were measured based on Alexa Fluor 680-conjugated BSA to label the circulating
plasma, while RBCs were identified based on their albumin-excluding negative label. The velocity
26

of RBCs (RBCV) was detected based on xt (line) scans performed in the central axis of the blood
vessels as described before (25, 36). Using the Leica LAS X software analysis tools (Version
3.7.4.23463), glomerular diameter, afferent (AA), and efferent (EA) arteriole internal diameters
(d) were measured. Single AA and EA blood flow (Q) was calculated based on the formula
Q=RBCV * (π * (d/2)
2
) and expressed in fL/msec.
2.3.6 Immunoblotting
Mouse kidney cortical tissue homogenates were prepared by manually dissecting the
kidney cortex and homogenizing for 2 minutes using Ultra-Turrax homogenizer (IKA Works, Inc.,
Wilmington, NC) in buffer containing 5% sorbitol, 25 mM histidine-imidazole (pH 7.5), 100 mM
Na2EDTA (pH 7.0) and 1X protease/phosphatase inhibitor cocktail (BD Bioscience, San Jose,
CA). Forty µg of protein was loaded in Any Kd pre-cast TGX gels (BioRad, Hercules, CA), stained
with Coomassie Blue dye (BioRad, Hercules, CA) and multiple bands were quantified to ensure
uniform loading of samples as described before (37, 38). For immunoblotting, 30-50 µg of protein
was loaded in wells of either 7.5% or Any Kd pre-cast TGX gels and then transferred onto a
polyvinylidene difluoride (PVDF) membrane (MilliporeSigma, Burlington, MA). Blots were
blocked for 30 minutes at room temperature using Odyssey Blocking Buffer (LI-COR Biosciences,
Lincoln, NE) followed by incubation in primary and secondary antibodies suspended in a 1% BSA
in 1X tris-buffered saline with 0.1% Tween (TBST) buffer sequentially overnight at 4ºC and for 1
hour at room temperature on a shaker, respectively. Primary antibodies and dilutions were as
follows: phospho p70S6 kinase (Cell Signaling Technology, CST9205, 1:500), renin (Anaspec
Inc., AS-54371, 1:1000), cyclooxygenase 2 (COX2) (Santa Cruz Biotechnology, sc-1745, 1:500),
microsomal prostaglandin E2 synthase 1 (mPGES1) (Cayman Chemical, 160140, 1:500), phospho
extracellular-signal regulated kinase 1/2 (ERK1/2) (CST4376, 1:500), total ERK1/2 (CST4696,
27

1:1000), phospho-p38 (CST9216, 1:1000), total p38 (CST8690, 1:1000) (all from Cell Signaling
Technology), CCN family member 1 (CCN1) (Santa Cruz Biotechnology, sc-13100, 1:500), CCN
family member 3 (CCN3) (Abcam, ab137677, 1:500), semaphorin 3C (Sema3C) (R&D Systems,
MAB1728, 1;500), C-X-C motif chemokine ligand 14 (CxCl14) (Novus Biologicals, NBP1-
31398, 1:500) and pappalysin 2 (Pappa2) (Invitrogen, PA5-21046). IRDye 680RD and IRDye
800CW secondary antibodies (LI-COR Biosciences) were all used at 1:5000 dilution. Blots were
then visualized using the Odyssey Infrared Imaging System (LI-COR Biosciences) and quantified
using Image Studio Software (LI-COR Biosciences).
2.3.7 Glomerular filtration rate measurements
For glomerular filtration rate (GFR) measurements, mice were injected retro-orbitally with
FITC-Sinistrin (7.5 mg/100 g body weight, MediBeacon, St. Louis, MO). GFR measurements
were performed using the MediBeacon Transdermal GFR Measurement System (MediBeacon) by
placing the sensor on the shaved skin for 90 minutes to detect fluorescence. GFR was then
calculated based on the decay kinetics of FITC-Sinistrin using MediBeacon Data Studio software
(MediBeacon).
2.3.8 Blood pressure measurements
Blood pressure (BP) was measured by tail-cuff plethysmography using a Visitech BP-2000
system (Visitech Systems, Apex, NC) as described before (30). All animals underwent a training
period of 5 days before the start of experimental measurements and data was expressed as an
average of measurements over 2 days.
2.3.9 Statistical methods
Data are expressed as mean ± SEM and were analyzed using Student’s t-test (between 2
groups) or one-way or two-way ANOVA (between multiple groups) with post-hoc comparison by
28

Tukey’s multiple comparisons test. Sample number n for each group is as indicated in the figure
legends. p < 0.05 was considered significant. Statistical analyses were performed using GraphPad
Prism 9.0 (GraphPad Software, Inc., San Diego, CA).

2.4 Results
2.4.1 Characterization of global protein synthesis in renal cell populations
Protein synthesis activity in renal cell populations is dynamic, reflecting functional and
structural differences between the cell types. In order to gain visual insights into the basal rate of
protein synthesis in the different cell types of the kidney, we developed SRY-Box transcription
factor 2 (Sox2)-tdTomato mice that feature tandem dimer Tomato (tdTomato) expression in all
renal cell populations (31). As seen in Fig. 2.1A, fluorescence imaging of frozen kidney tissue
sections of Sox2-tdTomato mice depicted constitutive and ubiquitous cytosolic expression of
tdTomato (in red) across the kidney cortex. A magnified fluorescence image of the kidney tissue
highlighted the differential expression of the tdTomato protein in specific cell types with the
highest intensity found in macula densa (MD) cells, followed by intercalated cells (ICs) and
podocytes as identified based on tissue morphology (Fig. 2.1B). To validate the identity of these
cell types, we performed immunofluorescence labeling with cell specific markers on frozen kidney
tissue sections. Co-localization of endogenous tdTomato expression (in red) with immunolabeling
of neuronal nitric oxide synthase (nNOS) for MD cells (Fig. 2.1C) (in cyan), p57 for podocytes
(Fig. 2.1D) (in cyan) and H-ATPase for ICs (Fig. 2.1E) (in cyan) confirmed these findings.
29

Amongst these three cell types, tdTomato protein expression appeared to be the highest in MD
cells, providing qualitative evidence of a high rate of synthesis in MD cells.
Next, to specifically determine the rate of overall protein synthesis in renal cell populations,
an O-propargyl-puromycin (OPP) incorporation-based fluorescence imaging approach was
implemented. As described previously (25, 35), this method relies on the development of the OPP
incorporated into the newly synthesized peptide strands using a fluorescent azide. Wildtype (WT)
Figure 2.1. Histological features of the Sox2-tdTomato mouse model. A: Representative fluorescence image of a SRY-
Box transcription factor 2 (Sox2)-tdTomato mouse kidney tissue section confirming the expression of tandem dimer
Tomato (tdTomato) in all kidney cell types (in red). Nuclei are labeled blue with DAPI. The specific cell types highlighted
are macula densa (MD) (solid white arrows), podocytes (dashed white arrows) and intercalated cells (ICs) (white
arrowheads). B: Magnified fluorescence image of a Sox2-tdTomato mouse kidney section focusing on a single glomerulus
with its MD cell plaque (solid white arrows) and podocytes (dashed white arrows). Neighboring ICs are also seen (white
arrowheads). Nuclei are labeled blue with DAPI. C-E: Representative immunofluorescence co-localization images for MD
cell marker neuronal nitric oxide synthase (nNOS) (in cyan) (C), podocyte marker p57 (in cyan) (D) and IC marker H-
ATPase (in cyan) (E) with endogenous tdTomato expression (in red). Note the intense tdTomato expression in these specific
cell types as compared to the other cell populations. G: glomerulus. Scale bars are 30 µm.
30

mice were given an intraperitoneal injection of either 20 mM OPP or dimethyl sulfoxide (DMSO)
as a vehicle control and the labeling was then developed on formalin-fixed paraffin-embedded
(FFPE) kidney tissue sections. To confirm the specificity of OPP in labeling nascent protein
strands, a subset of WT mice was also pre-treated with 16 mg/ml cycloheximide (CHX) (a potent
inhibitor of protein translation (33)) for 1 hour followed by OPP injection and this served as a
negative control. We did not observe any OPP labeling (FOPP) on FFPE kidney tissue sections of
mice injected with DMSO (0.47 ± 0.04, n = 4) (Fig. 2.2A, E) whereas strong OPP labeling (in red)
was observed in mice injected with OPP (0.94 ± 0.05, n = 6, p = 0.002) (Fig. 2.2B, E). Fluorescence
imaging of the kidney sections demonstrated the high level of protein synthesis in MD cells,
podocytes, and ICs with strong OPP labeling in the nuclear as well as cytoplasmic compartments.
Moreover, in mice pre-treated with CHX, OPP labeling was absent in almost all renal cell
populations, including MD cells (0.53 ± 0.03, n = 3, p = 0.0015) and podocytes (Fig. 2.2C, E).
However, faint OPP labeling was observed in the cytoplasm of a few scattered cells in distal
tubular segments (Fig. 2.2C). Finally, we treated WT mice with rapamycin (Rapa) to
pharmacologically inhibit mammalian target of rapamycin (mTOR) complex 1 (mTORC1), a
central regulator of cellular protein synthesis (21). Short-term Rapa treatment (8 mg/kg body
weight) for a period of 2 weeks (32) followed by OPP injection showed a marked reduction but
not a complete loss of OPP staining in MD cells (0.79 ± 0.08, n = 5) (Fig. 2.2D, E) as compared
to mice injected with OPP.
31

2.4.2 Generation and characterization of MD-mTOR
gof
mouse model
The central regulator of cellular protein synthesis is the mTOR signaling pathway via the
mTORC1 complex. We first verified the expression of several mTOR signaling proteins in the
human MD using immunohistochemistry data from the Human Protein Atlas (HPA) (39). As
depicted in Fig. 2.3A-D, the human MD plaque has a high level of expression of tuberous sclerosis
Figure 2.2. Quantitative visualization of protein synthesis activity in the kidney at the single-cell level using O-propargyl
puromycin-incorporation based fluorescence imaging. A-D: Representative fluorescence images of wildtype (WT) mouse
kidney sections without O-propargyl puromycin (OPP) (w/o OPP) (A), with OPP (w/ OPP) (in red) (B), with cycloheximide
(CHX) pre-treatment and OPP (in red) (C) and with Rapamycin (Rapa) treatment and OPP (in red) (D) with tissue
autofluorescence (in green) for morphological detail. Nuclei are labeled blue with DAPI. Note the strong OPP labeling (in red)
in the macula densa (MD) cell plaque (solid white arrows), podocytes (dashed white arrows) and intercalated cells (ICs) (white
arrowheads). E. Statistical summary of average OPP fluorescence intensity in the MD cell plaque (F OPP) normalized to red
blood cell (RBC) fluorescence intensity (F RBC) in WT mice w/o OPP (n = 4), w/ OPP (n = 6), with CHX pre-treatment and OPP
(n = 3) and Rapa treatment and OPP (n = 5). In each tissue section, multiple regions of interest (ROIs) were placed across 10
MD plaques and averaged. Data are expressed as mean ± SEM, **p <0.01, ***p< 0.001, one-way ANOVA with Dunnett’s
multiple comparisons test. G: glomerulus. Scale bars are 30 µm.
32

complex 2 (TSC2) (Fig. 2.3A), DEP domain containing mTOR interacting protein (Deptor) (Fig.
2.3B) along with ribosomal protein S6 kinase (RPS6K) (Fig. 2.3C) and eukaryotic translation
initiation factor 3C (eIF3C) (Fig. 2.3D). To specifically test the role of the mTOR signaling
pathway in regulating MD protein synthesis, we developed a MD-specific gain-of-function (gof)
of mTOR signaling mouse model using a Cre/lox driven approach. Using the classic MD-specific
marker nNOS, tamoxifen-inducible MD-mTOR
gof
mice were developed by intercrossing
nNOS/CreERT2 and TSC2/fl mice (29) on a C57BL6/J background. Further, these mice were
backcrossed with mTmG/fl mice to ensure MD-specific membrane targeted enhanced green
fluorescent protein (mG, eGFP) expression whereas all other cells in the kidney express membrane
targeted tdTomato (mT). Upon total tamoxifen induction (75 mg/kg tamoxifen via oral gavage
every alternate day for a total of 3 times), MD cells had targeted disruption of TSC2 (a negative
regulator of the mTORC1 complex (21)), along with membrane eGFP expression. Control MD-
GFP mice were generated by intercrossing nNOS/CreERT2 and mTmG/fl mice as described
before (5, 30), in which tamoxifen induction resulted in MD specific expression of membrane
eGFP. To validate the MD cell-specificity of altered TSC2 expression in MD-mTOR
gof
mice,
immunofluorescence labeling for TSC2 was performed on FFPE kidney tissue sections of control
and MD-mTOR
gof
mice post tamoxifen induction. Immunofluorescence imaging of kidney tissue
sections confirmed TSC2 expression (in red) in the MD of control mice (Fig. 2.3E) which was
specifically absent in the MD cell plaque in the MD-mTOR
gof
mice (Fig. 2.3F). To confirm the
upregulation of mTOR signaling in MD-mTOR
gof
mice, immunoblotting for phospho-p70S6
kinase (p-p70S6K) (an important downstream target of mTORC1(21)), was performed on kidney
cortex homogenates. As compared to control mice (126 ± 14, n = 7, p = 0.0140), MD-mTOR
gof
33

mice (200 ± 21, n = 7) had a significantly higher expression of p-p70S6K in the kidney cortex
(Fig. 2.3G).

Figure 2.3. Validation of the expression of mTOR signaling elements in human and control and MD-mTOR
gof
mouse
kidney. A-D: Human Protein Atlas (HPA) validation of immunohistochemistry labeling for tuberous sclerosis complex 2
(TSC2) (A), DEP domain containing mTOR interacting protein (Deptor) (B), ribosomal protein S6 kinase (RPS6K) (C) and
eukaryotic translation initiation factor 3C (eIF3C) (D) in human kidney. Magnified insets show labeling in the macula densa
(MD) (solid black arrows). Immunohistochemistry images for the above are available at: TSC2
(https://images.proteinatlas.org/30409/62646_A_9_5.jpg);Deptor(https://images.proteinatlas.org/23945/53320_A_7_5.jpg);
RPS6K(https://images.proteinatlas.org/2852/7862_A_8_5.jpg);eIF3C(https://images.proteinatlas.org/50112/117399_A_8_
5.jpg; E-F: Representative immunofluorescence images of mouse kidney tissue sections of control (E) and MD-mTOR
gof
(F)
mice with TSC2 labeling (in red) and tissue autofluorescence (in green) for morphological detail. Nuclei are labeled blue
with DAPI. Note the absence of TSC2 labeling in MD cells (solid white arrows) in Fig. 2.3F compared to in Fig. 2.3E. G:
Immunoblot for phospho-p70S6 kinase (p-p70S6K) in control and MD-mTOR
gof
kidney cortex homogenates (n = 7) with
statistical summary. Data are expressed as mean ± SEM, *p<0.05, Unpaired Student’s t-test. G: glomerulus. Scale bars are
30 µm.
34

2.4.3 Quantification of MD cell global protein synthesis in control and MD-mTOR
gof
mice
The OPP incorporation-based fluorescence imaging method (25) was utilized to measure
changes in MD protein synthesis activity in response to upregulated mTOR signaling and
physiological activation of MD cells using low salt (LS) diet (5). Control MD-GFP and MD-
mTOR
gof
mice were induced using tamoxifen at 14 weeks of age and after 4 weeks of induction,
mice were treated either with normal salt (NS) or LS diet for a period of 2 weeks. Mice were then
injected with OPP and labeling was developed on FFPE sections using a fluorescent azide. All
kidney tissue sections were imaged using the same confocal microscopy imaging settings and MD
cell plaques were identified based on their clearly discernible morphology (Fig. 2.4A-D). First, we
aimed to quantify changes in the rate of overall protein synthesis in response to upregulation of
mTOR signaling in MD cells. As compared to control mice on NS diet (0.94 ± 0.05, n = 6) (Fig.
2.4A, E), MD-mTOR
gof
mice on NS diet (1.30 ± 0.06, n = 8, p = 0.0004) (Fig. 2.4C, E) had
significantly increased protein synthesis activity, reflected by an increase in red fluorescence
intensity (FOPP) in the MD cell plaque. Furthermore, stimulation of the salt-sensing activity of MD
cells using LS diet increased the rate of protein synthesis by nearly 40% in control mice (1.40 ±
0.05 n = 6, p < 0.001) (Fig. 2.4B, E) and by nearly 20% in MD-mTOR
gof
mice (1.48 ± 0.05, n = 8)
(Fig. 2.4D, E) as compared to the respective mice on NS diet. The highest protein synthesis activity
was observed in the MD cell plaques of MD-mTOR
gof
mice on a LS diet (Fig. 2.4E). Additionally,
to confirm mTOR specificity of these responses, we treated control and MD-mTOR
gof
mice on LS
diet with Rapa (8 mg/kg body weight) for 2 weeks as described before (32) for pharmacological
inhibition of mTORC1 (21). Following Rapa treatment, we observed a significant reduction in MD
OPP labeling in both control mice on LS diet (LS: 1.40 ± 0.05, n = 6; LS+Rapa: 1.17 ± 0.04, n =
35

4; p = 0.0153), and in MD-mTOR
gof
mice on LS diet (LS: 1.48 ± 0.05, n = 8; LS+Rapa: 1.20 ±
0.02, n = 4, p = 0.0038) (data not shown).
Figure 2.4. Quantification of MD cell global protein synthesis in control and MD-mTOR
gof
mice using OPP-
incorporation based fluorescence imaging. A-D: Representative fluorescence images of mouse kidney tissue sections of
control mice on normal salt (NS) (A) or low salt (LS) (B) diet, MD-mTOR
gof
mice on NS (C) or LS (D) diet with O-
propargyl puromycin (OPP) labeling (in red) and tissue autofluorescence (in green) for morphological detail. Nuclei are
labeled blue with DAPI. Note the strong OPP labeling (in red) in the macula densa (MD) cell plaque (solid white arrows).
E: Statistical summary of average OPP fluorescence intensity in the MD cell plaque (F OPP) normalized to red blood cell
(RBC) fluorescence intensity (F RBC) in control mice on NS or LS diet (n = 6), and in MD-mTOR
gof
mice on NS or LS diet
(n = 8). In each tissue section, multiple regions of interest (ROIs) were placed across 10 MD plaques and averaged. Data
are expressed as mean ± SEM, ***p< 0.001, ****p <0.0001, one-way ANOVA with Tukey’s multiple comparisons test.
G: glomerulus. Scale bars are 25 µm.
36

2.4.4 Characterization of the autocrine effects of MD-mTOR signaling
In addition to playing a central role in protein synthesis and secretion, the mTOR signaling
pathway is known to regulate cell growth, proliferation, and autophagy (21). A recent publication
from our laboratory identified a unique feature of the MD cell microanatomy- the presence of a
dense network of major and minor processes named “maculapodia”, that are associated with a
secretory function and vesicular transport (5). To investigate the autocrine effect of upregulated
MD mTOR signaling, we quantified changes in the number of MD cells per plaque and the length
of the maculapodia (using mG expression to identify MD cells) in control MD-GFP and MD-
mTOR
gof
mice after tamoxifen induction. To assess both parameters, 25 µm thick frozen kidney
tissue sections were used to image the entire volume of the glomerulus via high resolution z-stacks.
Quantification of the length of maculapodia at the basal surface of MD cells showed a significant
increase in the length of the processes in MD-mTOR
gof
mice (5.91 ± 0.31 µm, n = 5) (Fig. 2.5B)
as compared to control mice (4.26 ± 0.23 µm, n = 8, p = 0.0011) (Fig. 2.5A, C). With respect to
the change in MD cell number, MD-mTOR
gof
mice had a significantly higher number of MD cells
per juxtaglomerular apparatus (JGA) area (12.00 ± 0.97 cells/area, n = 5) (Fig. 2.5E-F) compared
to control mice (9.20 ± 0.48 cells/area, n = 6, p = 0.0229) (Fig. 2.5D, F).
37

Figure 2.5. Autocrine and paracrine effects of upregulated MD mTOR signaling. A-B: Representative fluorescence
images of control (A) and MD-mTOR
gof
(B) mice with macula densa (MD) cells expressing enhanced green fluorescent
protein (eGFP) (in green) and all other cells expressing tandem dimer Tomato (tdTomato) (in red). Nuclei are labeled blue
with DAPI. Note the change in the length of maculapodia at the base of MD cells (solid white arrow). C: Statistical summary
of length of maculapodia in control (n = 8) and MD-mTOR
gof
(n = 5) mice. D-E: Representative fluorescence images of
control (D) and MD-mTOR
gof
(E) mice with MD cell plaque expressing eGFP (in green) and other cells expressing
tdTomato (in red). Nuclei are labeled blue with DAPI. Note the change in the number of MD cells per area (solid white
arrows). F: Statistical summary of number of MD cells per juxtaglomerular apparatus (JGA) area in control (n = 6) and
MD-mTOR
gof
(n = 5) mice. G: Statistical summary of systolic blood pressure (BP) of control and MD-mTOR
gof
mice on
either normal salt (NS) or low salt (LS) diet at baseline, 4 weeks (wk) post induction and 4 weeks post induction and 2
weeks of dietary treatment (n = 3-6). H: Statistical summary of glomerular filtration rate (GFR) in control (n = 10) and
MD-mTOR
gof
(n = 14) mice 4 weeks post induction. In each tissue section, 5-10 MD plaques were imaged via high
resolution z-stacks of the entire volume of the glomerulus and averaged. Data are expressed as mean ± SEM, *p<0.05, **p<
0.01, two-way ANOVA (between multiple groups) with Tukey’s multiple comparisons test, Unpaired Student’s t-test
(between 2 groups). G: glomerulus. Scale bars are 15 µm.
38

2.4.5 Blood pressure and kidney function in MD-mTOR
gof
mice
MD cells are chief regulators of glomerular filtration rate (GFR), renal blood flow (RBF),
and blood pressure (BP) via its effects on hemodynamics and the renin-angiotensin system (RAS)
(1). To elucidate the effect of upregulated MD mTOR signaling on kidney function, BP and GFR
of control and MD-mTOR
gof
mice were measured. As shown in Fig. 2.5G, no significant difference
was observed in the systolic BP of control and MD-mTOR
gof
mice at baseline, after induction as
well as after dietary salt treatment. Regarding kidney function, MD-mTOR
gof
mice had a
significantly elevated GFR after 4 weeks of induction (1981 ± 121 μL/min/100 g body weight, n
= 14) compared to control mice (1444 ± 99 μL/min/100 g body weight, n = 10, p = 0.0039) (Fig.
2.5H). No significant change was observed in the GFR after the LS dietary treatment in both mouse
strains (data not shown). In addition, no change was observed in the urinary albumin-to-creatinine
ratio (ACR) in control and MD-mTOR
gof
mice (data not shown).
2.4.6 Intravital imaging of glomerular hemodynamics in MD-mTOR
gof
mice
Using intravital multiphoton microscopy (MPM), changes in various structural and
functional parameters of glomerular hemodynamics at a single nephron level were quantified in
control MD-GFP and MD-mTOR
gof
mice after 7 days of tamoxifen induction. In both mouse
strains, MD cells expressed mG whereas all other cells in the kidney expressed mT. Intravital
imaging showed that MD-mTOR
gof
mice had a significantly enlarged glomerular tuft area (3021 ±
103 μm
2
, 27 glomeruli from n = 4 mice) as compared to control mice (2656 ± 87 μm
2
, 16 glomeruli
from n = 4 mice, p = 0.0198) (Fig. 2.6A). Next, we visualized the glomerular vascular pole region
at the base of the MD to quantify changes in the afferent arteriole (AA) and efferent arteriole (EA)
diameter and blood flow. MD-mTOR
gof
mice had significantly larger AA diameter (8.8 ± 0.4 μm,
21 glomeruli from n = 4 mice) (Fig. 2.6F, H) as compared to control mice (7.6 ± 0.3 μm, 16
glomeruli from n = 4 mice, p = 0.0345) (Fig. 2.6D, H). EA diameter measurements followed a
39

similar trend in glomeruli of MD-mTOR
gof
mice (6.6 ± 0.3 μm, 24 glomeruli from n = 4 mice)
(Fig. 2.6G, J) and control mice (5.3 ± 0.2 μm, 17 glomeruli from n = 4 mice, p = 0.0023) (Fig.
2.6E, J). Finally, xt (line) scans of the lumen of blood vessels were used to measure blood flow in
single AAs and EAs. AA blood flow was significantly higher in MD-mTOR
gof
mice (299 ± 76
fL/ms, 6 glomeruli from n = 4 mice) as compared to control mice (131 ± 9 fL/ms, 11 glomeruli
from n =4 mice, p = 0.0089) (Fig. 2.6I). Similar observations were made with respect to EA blood
flow in control mice (89 ± 10 fL/ms, 13 glomeruli from n = 4 mice, p = 0.0074) and MD-mTOR
gof
mice (163 ± 27 fL/ms, 8 glomeruli from n = 4 mice) (Fig. 2.6K). Additionally, we also quantified
changes in the diameter and blood flow within the glomerular capillaries (GCs). Similar to
observations made in the AA and EA, GC diameter was significantly elevated in MD-mTOR
gof

mice (7.0 ± 0.2 μm, 116 GCs from n = 4 mice) as compared to control mice (5.0 ± 0.1 μm, 80 GCs
from n = 4 mice, p < 0.0001) (Fig. 2.6B) along with significant increase in GC RBC velocity in
MD-mTOR
gof
mice (3.7 ± 0.4 μm/sec, 19 GCs from n = 4 mice) as compared to control mice (2.2
± 0.2 μm/sec, 17 glomerular capillaries from n = 4 mice, p = 0.0013) (Fig. 2.6C, and Supplemental
Video 1, https://figshare.com/s/245f48e344797eb22819).

40

Figure 2.6. Intravital imaging of glomerular hemodynamics in MD-mTOR
gof
mice. A-C: Statistical summary of
glomerular tuft area (A), glomerular capillary (GC) diameter (B) and GC red blood cell velocity (RBCV) (C) in control and
MD-mTOR
gof
mice (n = 4). D-G: Representative intravital multiphoton microscopy (MPM) images of glomeruli with their
afferent arteriole (AA) and efferent arteriole (EA) of control (D-E) and MD-mTOR
gof
(F-G) mice. Note the specificity of the
membrane targeted enhanced green fluorescent reporter (eGFP) expression (in green) to macula densa (MD) cells and
expression of membrane targeted tandem dimer Tomato (tdTomato) (in red) in all other renal cells. The circulating plasma is
labeled with Alexa Fluor 680-conjugated bovine serum albumin (BSA) (in grey). H-K: Statistical summary of AA diameter
(H), AA blood flow (I), EA diameter (J) and EA blood flow (K) in control and MD-mTOR
gof
mice (n = 4). Data are expressed
as mean ± SEM, *p<0.05, **p< 0.01, ****p<0.0001, Unpaired Student’s t-test. G: glomerulus. Scale bars are 30 µm.

41

2.4.7 Effects of MD mTOR signaling on renin expression
To determine the role of mTOR signaling in LS-driven physiological activation of MD
cells (5) and renin expression, immunofluorescence labeling for renin was performed on FFPE
kidney tissue sections of control and MD-mTOR
gof
mice treated with NS or LS diet for 2 weeks.
On NS diet, MD-mTOR
gof
mice displayed a significant increase in the number of renin
+
cells at
the vascular pole of the glomerulus (5.0 ± 0.3 cells/area, n = 8) (Fig. 2.7C, E) compared to control
mice (3.3 ± 0.2 cells/area, n = 6, p = 0.0018) (Fig. 2.7A, E). As expected, LS dietary treatment for
a period of 2 weeks resulted in a significant increase in the number of renin
+
cells in both control
mice (5.2 ± 0.2 cells/area, n = 6, p = 0.0018) (Fig. 2.7B, E) and MD-mTOR
gof
mice (6.3 ± 0.3
cells/area, n = 8, p = 0.0130) (Fig. 2.7D, E). However, the LS-induced increase in renin was
significantly higher in MD-mTOR
gof
mice as compared to control mice (p = 0.0445) (Fig. 2.7E).
Figure 2.7. Changes in renin expression in MD-mTOR
gof
mice. A-D: Representative maximum projection
immunofluorescence images of mouse kidney tissue sections of control mice on normal salt (NS) (A) or low salt (LS) (B)
diet, MD-mTOR
gof
mice on NS (C) or LS (D) diet with renin labeling (in red) and tissue autofluorescence (in green) for
morphological detail. Nuclei are labeled blue with DAPI. Macula densa (MD) cell plaque is highlighted (solid white arrows).
E: Statistical summary of the average number of renin
+
cells per juxtaglomerular apparatus (JGA) area in control (n = 6) and
MD-mTOR
gof
mice (n = 8) on either NS or LS diet. In each tissue section, 5-10 glomeruli were imaged via high resolution z-
stacks of the entire volume of the glomerulus and averaged. Data are expressed as mean ± SEM, *p<0.05, **p< 0.01, one-
way ANOVA with Tukey’s multiple comparisons test. G: glomerulus. Scale bars are 35 µm.
42

2.4.8 Changes in MD signaling in MD-mTOR
gof
mice
In order to address the effects of upregulated mTOR signaling on the elements of the classic
MD signaling pathway of renin secretion (cyclooxygenase 2 (COX2), microsomal prostaglandin
E2 synthase 1 (mPGES1), p38, and extracellular-signal regulated kinase 1/2 (ERK1/2) and renin)
(1) immunoblots for these proteins were quantified in the kidney cortex homogenates of control
and MD-mTOR
gof
mice (Fig. 2.8A). Mice were treated with tamoxifen at approximately 4 weeks
of age and tissue was harvested after 20 weeks of induction. Immunoblots for renin in kidney
cortex homogenates showed a trend to higher level of expression in MD-mTOR
gof
(616 ± 57, n =
7) compared to control (497 ± 17, n = 7, p = 0.0694) mice although it did not reach statistical
significance (Fig. 2.8A, B). COX2 expression was significantly higher in MD-mTOR
gof
mice
(3322 ± 203, n = 7) as compared to control mice (2689 ± 199, n = 7, p = 0.0228) (Fig. 2.8A, C).
Furthermore, immunoblots for ERK1/2 (Fig. 2.8A, D) and p38 (Fig. 2.8A, E) mitogen activated
protein kinases (MAPKs) showed no significant change in the ratio of phospho/total protein in
control and MD-mTOR
gof
mice. Similar to COX2, the expression of mPGES1 was significantly
higher in MD-mTOR
gof
mice (76 ± 4, n = 7) compared to control (56 ± 8, n = 7, p = 0.0379) (Fig.
2.8A, F).
Finally, since the overall rate of protein synthesis was significantly higher in MD-mTOR
gof

mice compared to control mice (Fig. 2.4E), we specifically probed for changes in the expression
levels of specific MD-enriched tissue remodeling and angiogenic peptides based on data from our
MD cell gene profile (unpublished data). These include two members of the CCN family- CCN1
and CCN3, C-X-C motif chemokine ligand 14 (CxCl14), semaphorin 3C (Sema3C), and
pappalysin 2 (Pappa2). CCN1 and CCN3 are both known to play an important role in angiogenesis
(40, 41), CxCl14 is a known chemokine (42), Sema3C is involved in axonal guidance (43) while
43

Pappa2 is a recently identified secreted protease highly expressed in MD cells (5, 44, 45).
Immunoblot analysis of expression of these proteins in kidney cortex homogenates showed a
significant increase all these proteins in MD-mTOR
gof
mice (CCN1: 2667 ± 192, CCN3: 9663 ±
574, CxCl14: 856 ± 74, Sema3C: 1293 ± 99, Pappa2: 700 ± 45, n = 7) in comparison to control
mice (CCN1: 1082 ± 174, p < 0.001, CCN3: 7127 ± 283, p = 0.0019, CxCl14: 646 ± 49, p =
0.0363, Sema3C: 934 ± 40, p = 0.0062, Pappa2: 584 ± 21, p = 0.0372, n = 7) (Fig. 2.8A, G-K).

44

Figure 2.8. Changes in MD signaling in MD-mTOR
gof
mice. A: Immunoblots for renin, cyclooxygenase 2 (COX2), total
extracellular-signal regulated kinase 1/2 (ERK1/2), phospho ERK1/2, total p38, phospho p38, microsomal prostaglandin E 2
synthase 1 (mPGES1), CCN family member 1 (CCN1), CCN family member 3 (CCN3), C-X-C motif chemokine ligand 14
(CxCl14), semaphorin 3C (Sema3C) and pappalysin 2 (Pappa2) in control and MD-mTOR
gof
mice kidney cortex homogenates
(n = 7). B-G: Statistical summary of immunoblot density for renin (B), COX2 (C), (phospho ERK1/2)/(total ERK1/2) (D),
(phospho p38)/(total p38) (E), mPGES1 (F), CCN1 (G), CCN3 (H), CxCl14 (I), Sema3C (J) and Pappa2 (K). Data are
expressed as mean ± SEM, *p<0.05, **p<0.01, ****p<0.0001, Unpaired Student’s t-test.
45

2.5 Discussion
The present study provided the first direct view of the high protein synthesis capacity of
the specialized cells of the macula densa (MD) and the autocrine and paracrine effects of
mammalian target of rapamycin (mTOR)-mediated upregulation of MD protein synthesis on
glomerular hemodynamics. A comprehensive experimental approach employed the use of novel
in vivo transgenic mouse models, intravital imaging techniques, and fluorescence assays along
with tissue, single-cell, and molecular tools including specific genetic and pharmacological
manipulation. Dual methodologies to visualize the basal rate of global protein synthesis at the
single-cell level, the SRY-Box transcription factor 2 (Sox2)- tdTomato mouse model, and the O-
propargyl puromycin (OPP) fluorescence assay confirmed the high rate of protein synthesis
activity in MD cells. Moreover, to understand the effect of increased MD protein synthesis on
renal function, we established and validated an MD-specific transgenic mouse model, MD-
mTOR
gof
mice to upregulate mTOR signaling specifically in MD cells. Upregulated MD mTOR
signaling and low salt diet (LS)-induced physiological activation of MD cells resulted in further
elevations of the rate of overall MD protein synthesis. Pharmacological inhibition with rapamycin
(Rapa) further confirmed the important regulatory role of mTOR signaling in MD cell protein
synthesis in both control and physiological activation conditions. Finally, quantification of the
traditional functions of MD cells in MD-mTOR
gof
versus control mice showed significant
elevations in glomerular hemodynamics, renal blood flow (RBF), and glomerular filtration rate
(GFR) along with an increase in the expression of renin and both classic and novel molecular
players in MD cell signaling including cyclooxygenase 2 (COX2) and pappalysin 2 (Pappa2). The
present study provided valuable new insight regarding the MD protein synthesis capacity and its
novel role in mediating cell-to-cell communications between the MD and surrounding cell
46

populations. The results from these experiments are consistent with our initial hypothesis regarding
the important role of the mTOR pathway in regulating MD protein synthesis activity as well as
enhancing the traditionally established function of MD cell in the kidney.
Since the identification of the MD cell plaque by Zimmerman and later works by
Goormaghtigh, the unique features of the polarized MD cell microanatomy- prominent primary
cilium and nucleus at the apical surface and numerous mitochondria, secretory organelles, and
cytoplasmic processes at the basal surface- have long been established using transmission electron
microscopy and conventional histology techniques (2, 17). These microanatomical features
suggested that MD cells are endowed with an extensive protein synthesis and secretory machinery
primarily located towards the basal surface of these cells. However, the complexity of the three-
dimensional space of the juxtaglomerular apparatus (JGA) and the relative inaccessibility of the
MD cell plaque have limited our ability to clarify the details of MD cell biological functions
including the rate of protein synthesis and the functional implications of these unique features of
MD cells. Recent advances in imaging techniques-especially intravital multiphoton microscopy
(MPM) (25, 27) - and the development of MD specific genetic mouse models have made it possible
to interrogate various aspects of MD cell biology in the intact living kidney (5).
Cellular protein synthesis is a tightly regulated process in order to maintain proteostasis;
disruption of the proteostatic networks between the endoplasmic reticulum (ER), mitochondria and
other protein synthetic organelles has been linked to the development of various
pathophysiological conditions including acute kidney injury, Fabry’s disease, and Alzheimer’s
disease (19, 22, 23). mTOR complex 1 (mTORC1) is the central regulator of cellular protein
synthesis via its effects on mRNA translation, ribosome biogenesis, and protein synthesis (21). To
visualize the basal level of global protein synthesis in the kidney in an unbiased fashion and at the
47

single-cell level, we used two distinct approaches-the Sox2-tdTomato mouse model and the OPP
fluorescence assay. In the newly-established Sox2-tdTomato genetic mouse model all renal cell
populations express the inert cytoplasmic red fluorescent protein tandem dimer Tomato
(tdTomato) under the control of the constitutively expressed Sox2/Cre (31) and the tdTomato
expression levels served as a surrogate marker for the direct qualitative visualization of the overall
rate of cellular protein synthesis. Interestingly, although the tdTomato expression was driven by a
constitutively active Cre, we observed differential expression of the fluorescent protein in specific
cell types in the kidney, with MD cells having the highest level of tdTomato followed by that in
collecting duct intercalated cells (ICs) and glomerular podocytes (Fig. 2.1A-E). These results
suggested that MD cells have the highest rate of global protein biosynthesis in the kidney cortex,
which is an interesting novel finding. To confirm these results, we next injected wildtype (WT)
mice with OPP in order to fluorescently label and specifically visualize nascent protein synthesis
(25, 35). In this histological imaging-based assay, the modified puromycin is added on to the newly
synthesized peptide strand, resulting in the metabolic labeling of proteins, making it possible to
quantify the rate of protein synthesis across different cell types. OPP labeling was observed in both
the nucleus and cytoplasm in all cells in the kidney, corresponding to the earliest synthesized
proteins (35). However, the labeling was stronger in MD cells, podocytes, and ICs, confirming the
observations from the Sox2-tdTomato mouse model (Fig. 2.2). Thus, both of these techniques
provided an unbiased spatial map of protein synthesis activity within the kidney cortex. As
expected, OPP labeling was completely abolished in MD cells in WT mice that were pre-treated
with cycloheximide (CHX), a potent inhibitor of protein translation (Fig. 2.2C) (33). Interestingly,
we observed that certain epithelial cells in the distal tubule (DT)-collecting duct, most likely ICs
based on their anatomy, retained a low level of OPP labeling even after CHX pre-treatment which
48

was not observed in either MD cells or podocytes (Fig. 2.2C). This observation suggests that in
addition to having a high level of protein synthesis, there may be a rapid rate of protein turnover
or secretion within MD cells in contrast to more protein storage in ICs that could be blocked with
a short-term pre-treatment with CHX. Finally, treatment with the mTORC1 inhibitor Rapa resulted
in an observable decrease in OPP labeling within the MD cell plaque. Interestingly, Rapa appeared
less effective in control conditions (Fig. 2.2D) compared to its significant inhibitory effects on MD
protein synthesis in the stimulated state (in MD-mTOR
gof
mice and LS diet), suggesting relatively
lower mTOR activity at baseline. Nevertheless, these results underscore the important role of the
mTOR signaling pathway in MD cells.
Immunohistochemistry labeling on human kidney tissue sections for several proteins in the
mTOR signaling cascade including tuberous sclerosis complex 2 (TSC2), and DEP domain
containing mTOR interacting protein (Deptor) ribosomal protein S6 kinase (RPS6K) and
eukaryotic translation initiation factor 3C (eIF3C) (Fig. 2.3A-D) from the Human Protein Atlas
(HPA) database (39) clearly suggested the high level of expression of these proteins in MD cells,
although the specificity of these immunolabeling results was not confirmed in the present study.
Moreover, we observed strong TSC2 labeling in MD cells on WT mouse kidney tissue sections
(Fig. 2.3E). Since TSC2 is a key negative regulator of mTORC1 (21), we used a neuronal nitric
oxide synthase (nNOS)/Cre driven approach (5) to disrupt TSC2 expression specifically in MD
cells and hence establish a new MD-mTOR gain-of-function (gof) signaling mouse model (MD-
mTOR
gof
) (29). The validation of the successful MD cell-specific genetic manipulation was
performed using immunofluorescence labeling for TSC2 on formalin-fixed paraffin-embedded
(FFPE) kidney tissue sections of control and MD-mTOR
gof
mice, which confirmed the presence
or absence of TSC2 expression in MD cells, respectively (Fig. 2.3E-F). Additionally, we also
49

found a significant increase in the expression of phospho-p70S6 kinase (p-p70S6K) in the kidney
cortex homogenates of MD-mTOR
gof
vs control mice (Fig. 2.3G), confirming the upregulation of
mTOR signaling activity (21).
Comparison of the rate of overall MD cell protein synthesis using the OPP assay in control
and MD-mTOR
gof
mice found that the upregulation of MD mTOR signaling resulted in a
significant increase in global MD protein synthesis (Fig. 2.4A, C). This increase in MD protein
synthesis was most likely due to the phosphorylation and activation of the ribosomal protein
p70S6K and the release of the inhibitory effect of the 4E binding protein 1 (4E-BP1) on the
eukaryotic translation initiation factor 4E (eIF4E) after the inhibitory effect of TSC2 on mTORC1
was genetically disrupted (21). Additionally, it is well-established that LS diet results in increased
activity of the p38 and ERK1/2 MAPKs and the physiological activation of MD cells (1, 5).
Reports from various eukaryotic model systems have demonstrated the extensive crosstalk
between the ERK1/2 and p38 mitogen activated protein kinases (MAPKs) and the mTOR signaling
pathway. For example, growth factor-derived activation of the MAPK pathway triggers the
activation of mTORC1 via its inhibition of the TSC1/TSC2 complex as well as by independently
activating p70S6K (46). Control mice treated with LS diet showed a discernible increase in OPP
labeling as compared to control mice on a normal salt (NS) diet (Fig. 2.4A-B). LS-based activation
and upregulation of MD mTOR signaling in MD-mTOR
gof
mice had an additive effect on MD
protein synthesis presumably due to the crosstalk between MAPK and mTOR intracellular
signaling cascades (Fig. 2.4D, E) since MAPKs are known to play a critical role in the traditional
function and LS activation of MD cells (1, 46). However, it is possible that other signaling
pathways (e.g., Ras-ERK) could be involved that activate mTOR downstream elements like
p70S6K independent of the mTORC1 complex.
50

Recent reports from our group (5) and others (3) have shown that the basal “maculapodia”
are likely associated with secretory function due to the presence of dense-core vesicles that contain
diverse cargo. Upregulated MD mTOR signaling resulted in a significant increase in the length of
these basal processes (Fig. 2.5A-C) and in MD cell number (Fig. 2.5D-F), supporting the proposed
role of maculapodia and MD cells in general in the regulation of paracrine communication to JGA
effector cells (5). While mTOR signaling is known to regulate cell proliferation and differentiation,
the exact mechanism responsible for the increased length of the maculapodia, and the identification
of the diverse cargo in secretory vesicles need further study.
The traditional role of MD cells as the tubular component of the JGA involves the
regulation of tubuloglomerular feedback (TGF) control of renal and glomerular hemodynamics
and renin release from the juxtaglomerular (JG) cells in response to alterations in tubular fluid salt
and flow rate (1, 6). To determine the overall functional consequences of altered MD cell protein
synthesis with respect to kidney function, we quantified changes in both GFR and RBF in response
to upregulated MD mTOR signaling. Using the MediBeacon transdermal sensor technique, we
measured changes in global GFR in control and in MD-mTOR
gof
mice 4 weeks after induction.
GFR was significantly higher in MD-mTOR
gof
mice with no observable change in systolic blood
pressure (BP) (Fig. 2.5G-H). In addition, the use of intravital multiphoton microscopy (MPM)
provided an unparalleled in-depth view of the otherwise inaccessible but critical vascular pole
region of the glomerulus along with high-resolution visualization of the afferent arteriole (AA)
and efferent arteriole (EA) that feed into the glomerulus (2, 27). The combination of using genetic
fluorescent reporters (eGFP) for easy identification of MD cells and quantitative intravital MPM
allowed us to uncover the robust elevation in RBF (including the first-time measurement of single
AA and EA blood flow in vivo) and vascular diameters in MD-mTOR
gof
versus control mice (Fig.
51

2.6 and Supplemental Video 1). Since the difference between the AA and EA RBF rate was greater
than the 20% filtration fraction (Fig. 2.6I, K) (36), these results further support the globally
measured elevated GFR in MD-mTOR
gof
mice (Fig. 2.5H).
Since MD cells play a critical role in the regulation of renin release from JG cells via the
release of prostaglandin E2 (PGE2) from the basal surface (1), we analyzed the changes in JG renin
cell number (and indirectly the JGA renin granule content) in response to upregulated mTOR
signaling and LS diet. Treatment of control and MD-mTOR
gof
mice with LS diet for 2 weeks
predictably resulted in a significant increase in renin expression at the glomerular vascular pole
(Fig. 2.7B, D-E). Interestingly, upregulation of MD mTOR signaling also resulted in a significant
elevation in renin expression (Fig. 2.7C, E) and this effect was even more pronounced in MD-
mTOR
gof
mice on a LS diet (Fig. 2.7D, E). These results suggest that the mTOR signaling pathway
has a combinatorial effect with the LS diet in regulating renin expression in the kidney cortex.
Moreover, the classic MD enzymes COX2 and microsomal prostaglandin E2 synthase 1 (mPGES1)
(1) also had significantly higher level of expression in MD-mTOR
gof
mice compared to control
mice (Fig. 2.8). Traditionally, these enzymes act downstream of the p38 and ERK1/2 MAPKs (1),
and the MAPK pathway is known to feed into the mTOR signaling cascade (46). However, since
we did not observe any changes in the expression or activity (phosphorylation) of p38 and ERK1/2
MAPKs, we propose that the MD mTOR pathway acts on COX2 and mPGES1 downstream of
MAPK signaling likely via selective regulation of protein synthesis.
Finally, our observation of increased protein synthesis in the MD led us to quantify changes
in specific MD-expressed proteins, both classic and novel. The increased expression of the classic
MD-specific proteins COX2 and mPGES1 (Fig. 2.8A) likely resulted in elevated PGE 2 synthesis
and release from MD cells which could explain, at least partially, the increased GFR, RBF, and
52

renin content found in MD-mTOR
gof
mice (Fig. 2.5H and Figs. 2.6-7). However, it is highly likely
that the increased expression of other MD cell proteins also contributed to the observed alterations
in glomerular hemodynamics and renin. Unpublished data from our laboratory shows high MD-
specific expression of several angiogenic- CCN family member 1 (CCN1), CCN family member
3 (CCN3), Pappa2, and tissue remodeling peptides- semaphorin 3C (Sema3C) and C-X-C motif
chemokine ligand 14 (CxCl14). Additionally, in a recent report from our laboratory, we have
shown the expression of Pappa2 in both human and mouse MD cells and its localization in likely
secretory type vesicles (5). In the present study, we observed a significant increase in CCN1,
CCN3, CxCl14, Sema3C, and Pappa2 expression in response to upregulated MD mTOR signaling
(Fig. 2.8A, G-K). These observations thus provide an initial clue regarding the nature of peptides
synthesized by MD cells. Future studies are needed to determine the functional significance of
these MD peptides and their regulation.
In summary, the present study discovered the high rate of MD protein synthesis by applying
direct imaging techniques with single-cell resolution and established the important regulatory role
of physiological activation and mTOR signaling in this process. This new feature of MD cells is a
novel component of the tubuloglomerular crosstalk and glomerular hemodynamic regulatory
functions of MD cells. Future work is needed to elucidate the nature and (patho)physiological role
of the specific proteins synthesized and likely secreted by MD cells, which will bring us closer to
the more complete understanding of this mysterious renal cell type.

53

Grants
This work was supported by US National Institutes of Health grants DK064324, DK123564, and
S10OD021833 to J.P-P.

Disclosures
J.P-P. and G.G. are co-founders of Macula Densa Cell LLC, a biotechnology company that
develops therapeutics to target macula densa cells for a regenerative treatment for chronic kidney
disease. Macula Densa Cell LLC has a patent entitled “Targeting macula densa cells as a new
therapeutic approach for kidney disease”. J. P-P. received consulting fees from Travere
Therapeutics and Eli Lilly &Co.

Author contributions
U.N.S., G.G., and J.P-P. conceived and designed research; U.N.S, G.G., A.I. and S.D. generated
materials and reagents, performed experiments and analyzed data; U.N.S. and G.G. prepared
figures, U.N.S. drafted manuscript; U.N.S., G.G., J.P-P. edited and revised manuscript; U.N.S,
G.G., A.I., S.D., and J.P-P. approved final version of manuscript.

Acknowledgments
U.N.S. was funded by predoctoral research fellowship 19PRE34380886 of the American Heart
Association.
54

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*
This chapter has been modified from my second author original research article “Gyarmati G, Shroff UN, Riquier-Brison A,
Desposito D, Izuhara A, Ahmadi N, Chen Y, Gill IS, and Peti-Peterdi J. Physiological activation of the nephron central command
drives kidney tissue remodeling and regeneration (in preparation)”. The methods appear in my first author book chapter “Shroff
UN, Schiessl IM, Gyarmati G, Riquier-Brison A, Peti-Peterdi J. Novel fluorescence techniques to quantitate renal cell biology.
Methods Cell Biol 154: 85-107, 2019”.

58

Chapter 3

Essential role of macula densa cell Wnt signaling in endogenous kidney tissue
remodeling
*

3.1 Abstract
The macula densa (MD), a plaque of 15-20 specialized cells, sits at the vascular pole of the
glomerulus and plays a traditional role in regulating renal hemodynamics and the renin-angiotensin
system (RAS). The canonical Wnt/β-catenin pathway is known to regulate several cellular
processes including embryonic development, cell proliferation, differentiation, and tissue
homeostasis. Since new results are emerging regarding MD-specific high expression of Wnt
signaling elements and target genes, the present study aimed to examine the role of the MD Wnt
signaling pathway in regulating glomerular structure and function. Newly generated transgenic
mouse models with MD-specific Wnt gain-of-function (gof) (MD-Wnt
gof
) and loss-of-function
(lof) (MD-Wnt
lof
), newly established fluorescence cell biology techniques, WntGFP reporter
mouse kidney tissue sections, and immunohistochemistry data from the Human Protein Atlas
(HPA) database showed that MD cells have the highest Wnt signaling activity in the renal cortex.
Changes in MD Wnt signaling activity resulted in several changes in glomerular architecture, with
an increase in glomerular size in MD-Wnt
gof
mice and smaller glomeruli in MD-Wnt
lof
mice.
Increased MD Wnt signaling significantly elevated glomerular filtration rate (GFR) and renin
expression with a corresponding decline in response to decreased MD Wnt signaling. Global rate

59

of MD protein synthesis, as well as the expression of MD-enriched proteins, was significantly
altered in both mouse models.
Finally, the up and down-regulation of MD Wnt signaling activity had robust effects on
glomerular tissue remodeling, resulting in changes in podocyte, endothelial cell, and endothelial
progenitor cell populations. The current study identified a dual role of the MD Wnt signaling
pathway in regulating both the traditional- renin, GFR- and non-traditional-tissue remodeling-
functions of MD cells in the kidney.
Keywords: macula densa, renin, glomerular filtration rate, remodeling

3.2 Introduction
Located at the end of the thick ascending limb (TAL) of the nephron, the macula densa
(MD) cell plaque consists of 20-25 highly specialized epithelial cells that are critical for the
regulation of body fluid and salt balance, glomerular hemodynamics via the tubuloglomerular
feedback (TGF) and renin release from the juxtaglomerular (JG) cells (1). The strategic
localization of MD cells at the glomerular vascular pole and its unique microanatomy- large apical
nucleus, basilar maculapodia (2), and densely packed secretory organelles at the basal surface- are
fundamental to mediate the chief sensory role of MD cells in the kidney (3). The traditional role
of MD cells is to respond to alterations in the tubular filtrate salt concentration, flow rate, and other
local tissue metabolites (1, 4). At the intracellular level, mitogen activated protein kinases
(MAPKs), mainly p38 and extracellular-signal regulated kinase 1/2 (ERK 1/2), along with
enzymes like microsomal prostaglandin E2 synthase 1 (mPGES1), cyclooxygenase 2 (COX2), and
neuronal nitric oxide synthase (nNOS), play an important role in regulating the traditional
60

functions of MD cells (5, 6). The autocoids synthesized as a result of the activation of these
enzymes - prostaglandin E2 (PGE2) and nitric oxide (NO)- have a significant paracrine effect on
the neighboring renal cell populations, mainly the JG renin cells (6, 7).
The canonical Wnt/β-catenin signaling pathway is known to play a critical role in various
developmental processes including the maintenance of stem cell populations and tissue
homeostasis (8). Specifically, the Wnt signaling pathway is an important regulator of epithelial
cell morphology, cell proliferation, differentiation, and homeostasis. The binding of the Wnt ligand
to its cognate Frizzled (Fzd)/low-density lipoprotein receptor-related protein 5/6 (LRP5/6) co-
receptor complex triggers the stabilization and translocation of β-catenin to the nucleus, where it
mediates the transcription of several Wnt target genes (9). Reports from other groups have shown
the major modulatory effect of NO and PGE2, two key MD cell-specific autocoids (6), on Wnt
signaling activity. Experimental data from human tumors and animal models have shown that NO
can negatively regulate the expression of Dickkopf 1 (DKK1), thus increasing Wnt signaling
activity (10) while data from hematopoietic stem cells has shown that PGE2 can enhance Wnt
signaling activity by regulating the degradation of β-catenin (11). Additionally, the Wnt signaling
pathway is also known to regulate cellular protein synthesis by activating the mammalian target of
rapamycin (mTOR) via inhibition of glycogen synthase kinase 3 (GSK3) (12).
While the role of PGE2 and NO in the traditional function of MD cells has been well-
established (6), the potential modulatory role of these autocoids with respect to the Wnt signaling
pathway in MD cells has not been studied. Recently published scRNAseq-based gene profile of
mouse MD cells (13), as well as our own bulk RNASeq-based MD cell transcriptional profiling
data (unpublished data), found high enrichment of Wnt10a and several Wnt target genes in MD
cells. However, the exact autocrine and paracrine roles of MD Wnt signaling have been unknown.
61

The development of MD-specific genetic mouse models and advancements in imaging techniques
have made it possible to overcome the technical impediments concerning the inaccessibility and
complexity of the juxtaglomerular apparatus (JGA), making it possible to discern the intricate
crosstalk between these intracellular signaling pathways (14, 15).
The current study aims to elucidate the role of the Wnt/β-catenin signaling pathway in
regulating MD cell function using newly established MD cell-specific gain/loss-of-function
(gof/lof) Wnt signaling mouse models. Furthermore, the study also explores the paracrine effect
of genetic manipulations of MD Wnt signaling on glomerular architecture and function.

3.3 Materials and Methods
3.3.1 Animals
All animal protocols were approved by the Institutional Animal Care and Use Committee
at the University of Southern California. Tamoxifen-inducible, conditional macula densa (MD)
cell-specific Wnt gain-of-function (gof) (MD-Wnt
gof
) mice on a mixed background were generated
by intercrossing nNOS/CreERT2 (2) and β-catenin/fl (exon 3 of β-catenin flanked by loxP sites)
(Jackson Laboratory, Bar Harbor, ME) (16) for MD-specific stabilization of β-catenin. Tamoxifen-
inducible, conditional MD cell-specific Wnt loss-of-function (lof) (MD-Wnt
lof
) mice on a
C57BL6/J background were similarly generated by intercrossing nNOS/CreERT2 (2) and β-
catenin/fl (exons 2-6 of β-catenin flanked by loxP sites) (Jackson Laboratory, Bar Harbor, ME)
(17) for MD-specific truncation of β-catenin. Both mouse strains were further backcrossed with
mTmG/fl reporter mice (Jackson Laboratory, Bar Harbor, ME) resulting in the expression of the
membrane targeted two-color fluorescent Cre-reporter tandem dimer Tomato (mT, tdTomato) and
enhanced green fluorescent protein (mG, eGFP) for MD-specific expression of eGFP after excision
62

(2). Tamoxifen-inducible, conditional MD-GFP mice on a C57BL6/J background were generated
by intercrossing nNOS/CreERT2 and mTmG/fl reporter mice (Jackson Laboratory, Bar Harbor,
ME) as described before (2). MD-Wnt
gof
:WntGFP and MD-Wnt
lof
:WntGFP mice were generated
by intercrossing MD-Wnt
gof
or MD-Wnt
lof
mice respectively with WntGFP reporter mice (Jackson
Laboratory, Bar Harbor, ME) (18).
3.3.2 Treatments
Mice were induced with tamoxifen (75 mg/kg body weight, Alfa Aesar, Haverhill, MA)
via oral gavage 3 times in total every alternate day. A subset of mice was treated with low salt (LS)
diet (TD 90228, Harlan Teklad, Madison, WI) for 2 weeks. O-propargyl-puromycin (OPP)
(Thermo Fisher Scientific, Waltham, MA) labeling was performed by injecting mice
intraperitoneally with 25 µl of 20 mM OPP dissolved in dimethyl sulfoxide (DMSO) for 1 hour
before tissue harvest and staining as described previously (15).
3.3.3 Tissue processing and immunofluorescence
Kidney tissue was harvested after anesthetizing mice with a combination of ketamine (100
mg/kg body weight) and xylazine (10 mg/kg body weight) and perfusing with ice-cold 1X
phosphate-buffered saline (PBS) and 4% paraformaldehyde (PFA) for 2 minutes using a peristaltic
pump. Following post-fixation with 4% PFA at room temperature, for frozen tissue blocks, tissue
was kept in 30% sucrose solution at 4ºC for cryoprotection and then embedded in Tissue-Tek
O.C.T. compound (Sakura Finetek, Torrance, CA) and flash frozen. Cryosections were cut 25 µm
thick, washed thrice with 1X PBS, and then mounted using DAPI-containing VectaShield
mounting media (Vector Laboratories Inc., Burlingame, CA). For paraffin tissue blocks, tissue was
dehydrated in increasing concentrations of ethanol followed by embedding in paraffin wax and
sectioned 8 µm thick. Immunofluorescence labeling for specific proteins was carried out by
63

incubating tissue sections in 0.5% Triton for 15 minutes followed by heat-induced antigen retrieval
with sodium citrate buffer (pH 6.0) for 12 minutes. Tissue sections were blocked using normal
donkey serum (1:20) for 30 minutes at room temperature to reduce non-specific labeling. Primary
and secondary antibodies were diluted in the blocking serum and were applied to the tissue sections
sequentially overnight at 4ºC and for 1 hour at room temperature, respectively, and mounted using
DAPI-containing VectaShield mounting medium (Vector Laboratories, Inc.). Primary antibodies
and dilutions were as follows-renin (Anaspec Inc., AS-54371, 1:100) and active β-catenin (ABC)
(MilliporeSigma, 05-665, 1:100). AlexaFluor 488 and 594-conjugated secondary antibodies
(Thermo Fisher Scientific) were used at 1:500 dilution. Tissue sections were imaged with a Leica
TCS SP8 DIVE (Leica Microsystems, Wetzlar, Germany) confocal/multiphoton laser scanning
microscope system as described previously (2).
3.3.4 OPP labeling and quantification
OPP labeling on tissue sections was developed using the AlexaFluor 594 Click-iT OPP
Protein Synthesis kit (Thermo Fisher Scientific) according to the manufacturer’s instructions (15).
All tissue sections were imaged using the identical laser power and imaging settings. To quantify
OPP intensity, multiple regions of interest (ROIs) were placed across the MD cell plaque (FOPP)
and then normalized to the red blood cell (RBC) fluorescence intensity (FRBC). Ten MD cell
plaques were quantified in each animal and data was represented as a mean value.
3.3.5 Immunoblotting
Mouse kidney cortex tissue homogenates were prepared by manually dissecting the kidney
cortex and homogenizing in buffer containing 5% sorbitol, 25 mM histidine-imidazole (pH 7.5),
100 mM Na2EDTA (pH 7.0), and 1X protease/phosphatase inhibitor cocktail (BD Bioscience, San
Jose, CA) for 2 minutes using Ultra-Turrax homogenizer (IKA Works, Inc., Wilmington, NC) on
64

ice. Forty µg of protein was loaded in Any Kd pre-cast TGX gels (Biorad, Hercules, CA), stained
with Coomassie Blue dye (Biorad) and multiple bands were quantified to ensure uniform loading
across the samples as described previously (19, 20). On average, 30-50 µg of protein was loaded
in either 7.5% or Any Kd pre-cast TGX gels (Biorad), transferred onto a polyvinylidene difluoride
(PVDF) membrane (MilliporeSigma, Burlington, CA), and then blocked using Odyssey Blocking
Buffer (LI-COR Biosciences, Lincoln, NE) for 30 minutes at room temperature on a shaker. Blots
were incubated sequentially in primary (overnight at 4ºC) and secondary antibodies (1 hour at
room temperature) diluted in a 1% bovine serum albumin (BSA) in 1X tris-buffered saline with
0.1% Tween (TBST). Primary antibodies and dilutions used were as follows- renin (Anaspec Inc.,
AS-54371, 1:1000), phospho extracellular-signal regulated kinase 1/2 (ERK1/2) (CST4376,
1:500), total ERK1/2 (CST4696, 1:1000) (both from Cell Signaling Technology) and CCN family
member 1 (CCN1) (Santa Cruz Biotechnology, sc-13100, 1:500). IRDye 680RD and IRDye
800CW secondary antibodies (LI-COR Biosciences) were all used at 1:5000 dilution. Blots were
then visualized using the Odyssey Infrared Imaging System (LI-COR Biosciences) and quantified
using Image Studio Software (LI-COR Biosciences).
3.3.6 Tissue clearing
Fresh kidney tissue slices were cut 500 µm thick and fixed with 4% PFA for 45 minutes at
room temperature, washed in 1X PBS, and blocked in a 2% SEA BLOCK (Thermo Fisher
Scientific) with 0.1% triton blocking solution for 1 hour in a 24-well µ-plate (ibidi GmbH,
Gräfelfing, Germany). Tissue slices were stained with primary and secondary antibodies diluted
in blocking solution in succession for 2 days at 4 ºC with continuous shaking. Primary antibodies
and dilutions were as follows- Wilms’ tumor 1 (WT1) (Abcam, ab89901, 1:1000) and Meis
homeobox 2 (Meis2) (MilliporeSigma, HPA003256, 1:200). AlexaFluor 488 and 594 conjugated
65

secondary antibodies (Thermo Fisher Scientific) were used at 1:500 dilution. Nuclei were labeled
using Hoechst (1:10000 in 1X PBS) for 1 hour at room temperature and then dehydrated using a
series of increasing concentrations of methanol (25%, 50%, and 75% methanol). Tissue clearing
was carried out by incubating tissue slices in a combination of 50:50 benzyl alcohol/benzyl
benzoate (1:2) (BABB): methanol solution and then stored in 100% BABB solution protected from
light (2, 15).
3.3.7 Glomerular filtration rate measurement
Glomerular filtration rate (GFR) measurements were performed by injecting mice retro-
orbitally with FITC-Sinistrin dye (7.5 mg/100 g body weight, MediBeacon, St. Louis, MO). Mice
were shaved using depilatory cream and the MediBeacon Transdermal GFR sensor (MediBeacon)
was placed firmly on the shaved skin to measure the decay kinetics of the FITC-Sinistrin dye. GFR
was calculated using the MediBeacon Data Studio software (MediBeacon).
3.3.8 Statistical methods
Data are expressed as mean ± SEM. Statistical analyses were performed using unpaired
Student’s t- test (between 2 groups) or using one-way ANOVA (for multiple groups) with post-
hoc analysis using Tukey’s multiple comparison test. Sample size n for each group is as indicated
in figure legends. p < 0.05 was considered statistically significant. Statistical analyses were
performed using GraphPad Prism 9.0 software (GraphPad Software, Inc., San Diego, CA).

66

3.4 Results
3.4.1 Characterization of Wnt/β-catenin signaling activity in macula densa cells
The Wnt/β-catenin signaling pathway plays a critical role in maintaining epithelial cell
morphology, proliferation, and gene expression (8). We first used a commonly available genetic
mouse model- the WntGFP reporter mouse model in order to determine the level of Wnt signaling
activity in macula densa (MD) cells in the kidney cortex. In these reporter mice, the green
fluorescent protein (GFP) expression in the nucleus is driven by the level of Wnt signaling activity,
thus allowing for high-resolution single cell visualization of Wnt signaling activity across different
cell types (18). Using this mouse model, we characterized the level of Wnt signaling activity in
the kidney cortex on thick frozen tissue sections via high resolution z-stack confocal imaging. As
observed in Fig. 3.1A, as compared to the neighboring cell populations, the entire MD cell plaque
displayed intense GFP expression, suggesting the highest Wnt signaling activity in these cells.
Very few cells within the glomerulus and proximal tubule (PT) were positive for GFP expression
while on average GFP expression was higher in the distal tubule (DT) compared to PT segments.
Additionally, we also carried out immunofluorescence labeling for active-β-catenin (ABC), the
final downstream effector of the Wnt signaling pathway (9), on wildtype (WT) mouse kidney
67

tissue sections. Qualitative fluorescence imaging showed high expression of ABC (in green) in
MD cells, within both the cytoplasmic and nuclear compartments (Fig. 3.1B). Finally, we also
used the Human Protein Atlas (HPA) database (21) to validate the high level of Wnt signaling
activity in the human MD. Immunohistochemistry data from the HPA database showed that MD
Figure 3.1. Wnt/β-catenin signaling activity in macula densa cells. A: Representative fluorescence image of a WntGFP
reporter mouse kidney tissue section depicting Wnt signaling activity (in green) in the kidney cortex. Note the intense green
fluorescent protein (GFP) expression in macula densa (MD) cell plaque (solid white arrows) compared to other kidney cell
types. B: Representative immunofluorescence image of wildtype (WT) mouse kidney tissue section with active β-catenin
(ABC) labeling (in green) in MD cells (solid white arrows). C-E: Immunohistochemistry labeling for glycogen synthase
kinase 3β (GSK3β) (C), T-cell factor 4 (TCF4) (D) and lymphoid-enhancer binding factor 1 (LEF1) (E) from the Human
Protein Atlas (HPA) database. Magnified insets show labeling in the MD (solid black arrows). Immunohistochemistry images
for the above are available at:GSK3β(https://images.proteinatlas.org/16263/36255_A_7_5.jpg);TCF4
(https://images.proteinatlas.org/25958/55485_A_7_5.jpg); LEF1 (https://images.proteinatlas.org/2087/7387_A_9_5.jpg); G:
glomerulus, PT: proximal tubule, DT: distal tubule. Scale bars are 30 µm.
Image credit:
Human Protein Atlas
Image credit:
Human Protein Atlas
Image credit:
Human Protein Atlas
68

cells had strong immunolabeling for several Wnt signaling cascade elements compared to other
renal cell types including glycogen synthase kinase 3β (GSK3β) (Fig. 3.1C), T-cell factor 4 (TCF4)
(Fig. 3.1D) and lymphoid-enhancer binding factor 1 (LEF1) (Fig. 3.1E).
3.4.2 Generation and validation of MD-Wnt
lof
and MD-Wnt
gof
mouse models
To determine the role of the Wnt signaling pathway in MD cells, we developed MD-
specific gain-of-function (gof) and loss-of-function (lof) Wnt signaling inducible mouse models-
MD-Wnt
gof
and MD-Wnt
lof
respectively- using a Cre/lox driven approach. For both mouse models,
we used a nNOS/Cre mouse since expression of neuronal nitric oxide synthase (nNOS) is specific
to the MD in the kidney (2) (Fig. 3.2A). In the MD-Wnt
gof
mice, upon induction, there is MD-
specific stabilization of β-catenin and thus, upregulated Wnt signaling (16) while in the MD-Wnt
lof

mice, MD cells express a truncated β-catenin, resulting in downregulation of Wnt signaling (17).
To confirm these alterations in MD Wnt signaling activity, we further backcrossed these
mice with the WntGFP reporter mouse model (18). Fluorescence images of kidney tissue from the
MD-Wnt
gof
:WntGFP reporter mice (Fig. 3.2D) showed that MD cells in these mice had a higher
GFP expression as compared to control WntGFP mice (Fig. 3.2B). Moreover, in the MD-
Wnt
lof
:WntGFP mice (Fig. 3.2C), GFP expression in the MD cell plaque was nearly absent with
no observable change in any other cell type in the kidney.
69

3.4.3 Morphological and functional characterization of MD-Wnt
lof
and MD-Wnt
gof
mouse
models
Morphological phenotyping of kidney tissue of the MD-Wnt
lof
and MD-Wnt
gof
mice was
performed via periodic acid Schiff (PAS) staining. Compared to control WT mice (Fig. 3.3A, D),
MD-Wnt
lof
mice had hypomorphic and sclerotic glomeruli with smaller glomerular diameter (Fig.
3.3B, D), while MD-Wnt
gof
mice featured enlarged and hypercellular glomeruli (Fig. 3.3 C-D). As
quantified in Fig. 3.3E-F, no significant change was observed in either the kidney weight
normalized to body weight or the body weight of all mouse strains.
Figure 3.2. Validation of MD-Wnt
lof
and MD-Wnt
gof
mouse models. A: Schematic of the Cre/lox based breeding and
genetic strategy to generate MD-Wnt
gof
and MD-Wnt
lof
mice. B-D: Representative fluorescence images of control WntGFP
(B), MD-Wnt
lof
:WntGFP (C) and MD-Wnt
gof
:WntGFP (D) mouse kidney tissue sections with endogenous WntGFP expression
(in green) and differential interference contrast (DIC) (in grey) for morphological detail. Note the complete loss of green
fluorescent protein (GFP) expression in MD-Wnt
lof
mice in macula densa (MD) cell plaque (solid white arrows) (Fig. 3.2C)
and intense green fluorescent protein (GFP) expression in MD-Wnt
gof
mice (Fig. 3.2D). G: glomerulus. Scale bars are 25 µm.
70

To determine the effect of altered MD Wnt signaling on renal function, we measured the
glomerular filtration rate (GFR) and systolic blood pressure (BP) in these mice after 4 weeks of
tamoxifen induction. As shown in Fig. 3.3G, we did not observe any significant difference in the
systolic BP of both MD-Wnt
lof
and MD-Wnt
gof
mice as compared to the control mice after 4 weeks
of induction. To measure changes in the GFR, we used the transdermal MediBeacon sensor
technique and tracked the fluorescence decay kinetics of the injected FITC-Sinistrin dye. As
compared to the control mice (1559 ± 69 µL/min/100 g body weight, n = 10, p = 0.0160), we
observed a significant increase in GFR in MD-Wnt
gof
mice (2080 ± 202 µL/min/100 g body weight,
n = 5) (Fig. 3.3H). In contrast, GFR was significantly decreased in the MD-Wnt
lof
mice (1173 ±
132 µL/min/100 g body weight, n = 7) compared to the control mice (1559 ± 69 µL/min/100 g
body weight, n = 10, p = 0.0486) (Fig. 3.3H). Finally, we also measured changes in albuminuria
71

of these mice after tamoxifen induction; no significant changes were observed in either strains of
mice (data not shown).
Figure 3.3. Morphological and functional characterization of MD-Wnt
lof
and MD-Wnt
gof
mouse models. A-C:
Representative images of mouse kidney tissue sections of control (A), MD-Wnt
lof
(B) and MD-Wnt
gof
mice

(C) with periodic
acid Schiff (PAS) staining. Note the reduced glomerular size in MD-Wnt
lof
kidneys compared to control, and the enlarged
and hypercellular glomeruli in MD-Wnt
gof
mouse kidneys. D-E: Statistical summary of the glomerular diameter (D),
changes in kidney weight normalized to body weight (E), and body weight (F) in control (n = 8), MD-Wnt
lof
(n = 4)

and
MD-Wnt
gof
mice (n = 5). G: Statistical summary of systolic blood pressure (BP) of control, MD-Wnt
lof
and MD-Wnt
gof
mice
4 weeks after induction (n = 7-10). H: Statistical summary of glomerular filtration rate (GFR) in control (n = 10), MD-
Wnt
lof
(n = 7) and MD-Wnt
gof
(n = 5) mice. In each tissue section, diameter of 10-20 glomeruli was measured via high
resolution z-stacks of the entire volume of the glomerulus. Data are expressed as mean ± SEM, *p<0.05, ****p< 0.0001,
one-way ANOVA with Dunnett’s multiple comparisons test.
72

3.4.4 Paracrine effect of MD Wnt signaling
As a part of the juxtaglomerular apparatus (JGA), MD cells play a key role in mediating
renin release from the juxtaglomerular (JG) cells in response to a lower tubular filtrate salt
concentration (6). We first aimed to quantify changes in renin expression in response to altered
MD Wnt signaling and physiological activation via low salt (LS) diet (2) by carrying out
immunofluorescence labeling for renin on formalin-fixed paraffin-embedded (FFPE) kidney tissue
sections. As quantified in Fig. 3.4G, compared to control mice on a normal salt (NS) diet (3.3 ±
0.2 cells/area, n = 6, p = 0.0076) (Fig. 3.4A), MD-Wnt
gof
mice on a NS diet (6.4 ± 0.3 cells/area, n
= 3) (Fig. 3.4E) had a significant elevation in the number of renin
+
cells at the glomerular vascular
pole. No significant change could be detected in the granular renin content in the MD-Wnt
lof
mice
on NS diet. LS treatment resulted in elevated renin expression across all groups of mice (Fig. 3.4B,
D, F). Immunoblot quantification of renin expression in kidney cortex homogenates showed a
significant decrease in renin expression in MD-Wnt
lof
mice (437 ± 26, n = 4, p = 0.0443) as
compared to control mice (516 ± 18, n = 4) (Fig. 3.4H) whereas there was a significant increase in
renin expression in MD-Wnt
gof
mice (739 ± 35, n = 5, p = 0.0185) as compared to control mice
(587 ± 34, n = 4) (Fig. 3.4I).We did not observe any change in the ratio of phospho-to-total
extracellular-signal regulated kinase 1/2 (ERK1/2) in either MD-Wnt
lof
mice (Fig. 3.4J) or MD-
Wnt
gof
(Fig. 3.4K).

73

Figure 3.4. Paracrine effect of MD Wnt signaling on renin expression. A-F: Representative maximum projection
immunofluorescence images of mouse kidney tissue z-sections of control mice on normal salt (NS) (A) or low salt (LS)
(B) diet, MD-Wnt
lof
mice on NS (C) or LS (D) diet and MD-Wnt
gof
mice on NS (E) or LS (F) diet with renin labeling (in
red) and tissue autofluorescence (in green) for morphological detail. Nuclei are labeled blue with DAPI. Macula densa
(MD) cell plaque is highlighted (solid white arrows). G: Statistical summary of the average number of renin
+
cells/area in
control (n = 6), MD-Wnt
lof
(n = 4) and MD-Wnt
gof
(n = 3-4) mice on either NS or LS diet. H-I: Immunoblots for renin in
control (n = 4) and MD-Wnt
lof
(n = 4) (H) or MD-Wnt
gof
(n = 5) (I) mice kidney cortex homogenates with statistical
summary. J-K: Immunoblots for phospho-extracellular-signal regulated kinase 1/2 (ERK1/2) and total ERK1/2 in control
(n = 4) and MD-Wnt
lof
(n = 4) (J) or MD-Wnt
gof
(n = 5) (K) mice with statistical summary. In each tissue section, 5-10
glomeruli were imaged via high resolution z-stacks of the entire volume of the glomerulus and averaged. Data are expressed
as mean ± SEM, *p<0.05, **p< 0.01, one-way ANOVA (between multiple groups) with Tukey’s multiple comparisons
test, Unpaired Student’s t-test (between 2 groups). G: glomerulus. Scale bars are 30 µm.
74

3.4.5 Regulation of MD cell protein synthesis by MD Wnt signaling
Cellular protein synthesis is critical in determining the fate and function of a specific cell
type, and reports from other groups have established the role of the Wnt signaling pathway in
regulating cellular protein synthesis via its effect on the mammalian target of rapamycin (mTOR)
signaling pathway (12). In order to determine the regulatory effect of the Wnt signaling pathway
on MD protein synthesis activity, we used the newly-established O-propargyl-puromycin (OPP)-
incorporation based fluorescence imaging technique (15, 22) and quantified the rate of global
protein synthesis in MD cells. Mice (14 week old) were induced with tamoxifen for a period of 4
weeks followed by treatment with either NS or LS diet for 2 weeks. OPP labeling was developed
on FFPE sections using a fluorescent azide and rate of protein synthesis was calculated based on
the intensity of the OPP labeling (FOPP). As compared to control mice on a NS diet (0.94 ± 0.05, n
= 6, p = 0.0007) (Fig. 3.5A), MD-Wnt
gof
mice on a NS diet (1.38 ± 0.05, n = 4) (Fig. 3.5E) had a
significantly higher rate of global protein synthesis within the MD cell plaque with no change in
the MD-Wnt
lof
mice on a NS diet (1.01 ± 0.05, n = 4) (Fig. 3.5C). Physiological activation of MD
cells with LS diet resulted in a significant increase in OPP fluorescence intensity in control mice
(1.40 ± 0.05, n = 6, p < 0.0001) (Fig. 3.5B) and tended to increase in MD-Wnt
lof
mice but did not
reach statistical significance (1.31 ± 0.07, n = 4, p = 0.0617) (Fig. 3.5D). Highest rate of protein
synthesis was observed in MD-Wnt
gof
mice on a LS diet (1.59 ± 0.10, n = 4) as reflected by the
intense OPP labeling (Fig. 3.5F).
Gene profile analysis of mouse MD cells via bulk RNASeq demonstrated high level of
enrichment of several growth factors, angiogenic factors, and tissue remodeling peptides in MD
75

Figure 3.5. Quantification of MD cell protein synthesis in response to altered MD Wnt signaling using O-propargyl-
puromycin-incorporation based fluorescence imaging. A-F: Representative fluorescence images of mouse kidney tissue
sections of control mice on normal salt (NS) (A) or low salt (LS) (B) diet, MD-Wnt
lof
mice on NS (C) or LS (D) diet and
MD-Wnt
gof
mice on NS (E) or LS (F) diet with O-propargyl-puromycin (OPP) labeling (in red) and tissue autofluorescence
(in green) for morphological detail. Nuclei are labeled blue with DAPI. Note the strong OPP labeling in the macula densa
(MD) cells (solid white arrows). G: Statistical summary of the average OPP fluorescence intensities in the MD cell plaque
(F OPP) normalized to red blood cell (RBC) fluorescence intensity (F RBC) in control mice on NS or LS diet (n = 6) and MD-
Wnt
lof
or MD-Wnt
gof
mice on NS or LS diet (n = 4). H-I: Immunoblot for the angiogenic modulator protein CCN family
member 1 (CCN1) in control (n = 4) and MD-Wnt
lof
(n = 4) (H) or MD-Wnt
gof
(n = 5) (I) mice with statistical summary. In
each tissue section, multiple regions of interest were placed across 10 MD plaques and averaged. Data are expressed as
mean ± SEM, *p<0.05, **p< 0.01, ***p< 0.001, ****p< 0.0001, one-way ANOVA (between multiple groups) with Šidák’s
multiple comparisons test, Unpaired Student’s t-test (between 2 groups). G: glomerulus. Scale bars are 20 µm.
76

cells as compared to other distal tubular cells. MD-enriched angiogenic factors include CCN
family member 1 (CCN1), pappalysin 2 (Pappa2), semaphorin 3C (Sema3C), C-X-C motif
chemokine ligand 14 (CxCl14), and CCN family member 3 (CCN3) (unpublished data). Moreover,
data from HPA database (21) corroborated the high level of expression of these proteins in the
human MD (data not shown). As a proof-of-concept, we quantified changes in the level of
expression of CCN1, a known Wnt-target gene and a potent regulator of angiogenesis (23, 24), in
response to altered MD Wnt signaling. Immunoblot analysis of kidney cortex homogenates for
CCN1 expression showed a significant decrease in CCN1 expression in the MD-Wnt
lof
mice (1067
± 32, n = 4) compared to control mice (1291 ± 49, n = 4, p = 0.0082) (Fig. 3.5H). The opposite
trend was observed in MD- Wnt
gof
mice (869 ± 105, n = 5, p = 0.1060) compared to control mice
(579 ± 116, n = 4) (Fig. 3.5I).
3.4.6 Tissue remodeling activity of MD Wnt signaling
To determine the effect of MD Wnt signaling activity on neighboring renal cell
populations-particularly podocytes and endothelial cells-we carried out immunofluorescence
labeling and three-dimensional imaging with cell specific markers in cleared kidney tissue. First,
we quantified changes in the number of podocytes within the glomerulus using Wilms’ tumor 1
(WT1) as a marker. As compared to control mice (85 ± 2 cells/glomerulus, n = 7, p < 0.0001),
MD-Wnt
lof
mice had a significantly lower number of WT1
+
cells (56 ± 1 cells/glomerulus, n = 4).
On the other hand, MD-Wnt
gof
mice (92 ± 2 cells/glomerulus, n = 4) showed a significant increase
in the number of WT1
+
podocytes compared to control (85 ± 2 cells/glomerulus, n = 7, p = 0.0303)
(Fig. 3.6A, D).
77

Similarly, using Meis homeobox 2 (Meis2) as a marker for endothelial cells, we observed
a significant increase in the number of glomerular endothelial cells in MD-Wnt
gof
mice (85 ± 4, n
Figure 3.6. Tissue remodeling effect of MD Wnt signaling. A-B: Representative fluorescence images of mouse kidney
tissue sections of control, MD-Wnt
lof
and MD-Wnt
gof
mice with Wilms’ tumor 1 (WT1) labeling (in red) (A), Meis
homeobox 2 (Meis2) (in red) (B) and tissue autofluorescence (in green) for morphological detail. Nuclei are labeled blue
with DAPI. Macula densa (MD) cell plaque is highlighted (solid white arrows). C: Representative fluorescence images of
mouse kidney tissue sections of control, MD-Wnt
lof
and MD-Wnt
gof
mice with CD34 labeling (in green) and tissue
autofluorescence (in yellow) for morphological detail. Nuclei are labeled blue with DAPI. MD cells are highlighted (solid
white arrows). D-F: Statistical summary of the average number of WT1
+
cells/glomerulus

(D), Meis2
+
cells/glomerulus (E)
and CD34
+
cells/area in control (n = 7-8), MD-Wnt
lof
(n = 4) and MD-Wnt
gof
mice

(n = 4). Data are expressed as mean ±
SEM, *p<0.05, **p<0.01, ****p<0.001 one-way ANOVA with Dunnett’s multiple comparisons test. G: glomerulus. Scale
bars are 25 µm.

78

= 4) compared to control mice (71 ± 2, n = 8, p = 0.0043). In contrast the downregulation of MD
Wnt signaling in MD-Wnt
lof
mice (57 ± 3, n = 4) resulted in a significant decrease in Meis2
+
cells
as compared to control mice (71 ± 2, n = 8, p = 0.0069) (Fig. 3.6B, E).
Finally, we used CD34 as a marker for endothelial progenitor cells (25) for
immunofluorescence labeling on kidney tissue sections. The number of glomerular CD34
+

endothelial cells was significantly higher in MD-Wnt
gof
mice (16 ± 2 cells/area, 13 glomeruli from
n = 4 mice) as compared to control mice (2 ± 0.4 cells/area, 13 glomeruli from n = 4 mice, p <
0.0001) whereas no change was observed in MD-Wnt
lof
mice (Fig. 3.6C, F). Interestingly, the
CD34
+
staining in MD-Wnt
gof
mouse kidney tissue sections was predominantly localized at the
vascular entrance at the base of the MD cells (Fig. 3.6C).

3.5. Discussion
Using macula densa (MD)-specific genetic mouse models and state-of-the-art imaging
techniques, the current study explores the novel role of the Wnt/β-catenin signaling pathway in
regulating MD cell function and its paracrine effect on glomerular architecture and renal function.
The focus on the role of Wnt/β-catenin signaling pathway was driven by several observations with
regard to the level of Wnt signaling activity in MD cells; the WntGFP reporter mouse model (18)
showed highest green fluorescent protein (GFP) expression within the MD cell plaque along with
elevated expression of several proteins involved in this pathway (13). In order to address the
autocrine and paracrine regulatory roles of the MD Wnt signaling pathway, we first generated and
validated MD-specific genetic mouse models to either upregulate (MD-Wnt
gof
) or downregulate
(MD-Wnt
lof
) Wnt signaling activity using a Cre/lox-based approach to alter the expression of β-
79

catenin, the downstream target of Wnt signaling (9). Using these genetic mouse models, we
quantified the effect of the MD Wnt signaling pathway on renal function, specifically glomerular
filtration rate (GFR). Additionally, using a O-propargyl-puromycin (OPP) incorporation-based
fluorescence imaging approach (15, 22), we were able to quantify changes in the rate of global
protein synthesis in MD cells. With respect to the traditional functions of MD cells in the kidney,
we observed a robust effect of MD Wnt signaling on renin expression. Finally, we also explored
the non-traditional role of MD cells in mediating endogenous glomerular tissue remodeling. We
observed significant changes in the glomerular architecture-specifically increased podocyte and
endothelial cell numbers-in response to upregulated MD Wnt signaling. The present study
established the novel important roles of Wnt/β-catenin signaling in MD cells with respect to
cellular protein synthesis (autocrine role) and the regulation of renin and glomerular cell types and
tissue remodeling (paracrine role).
The fundamental role of the Wnt signaling pathway in the development of the various
organ systems including the kidney, maintenance of stem cell populations, and regulation of
epithelial cell morphology and function has long been established (8). For example, Wnt9b and
Wnt4 play a critical role in the induction of renal vesicles from the metanephric mesenchyme (26).
However, the role of this signaling pathway in the highly specialized epithelial MD cells, has been
unknown, partly due to the complex architecture of MD cells and technical limitations in
specifically targeting MD cells. Advances made in the development of MD-specific genetic mouse
models (2) and imaging techniques (15) have now made it possible to interrogate the role of a
particular signaling cascade within these cells.
The Wnt/β-catenin signaling pathway controls gene expression via the T-cell factor
(TCF)/lymphoid-enhancer binding factor (LEF) family of transcription factors (9). Moreover, Wnt
80

signaling mediated phosphorylation of glycogen synthase kinase 3 (GSK3) promotes protein
translation via the mammalian target of rapamycin (mTOR) signaling pathway (12). Thus, the Wnt
pathway has a dual role in controlling both gene expression and protein synthesis. To establish the
basal level of Wnt signaling in MD cells, we first made use of a fluorescent reporter mouse model-
the WntGFP mouse model. In these mice, the intensity of GFP expression acts as a readout of the
level of Wnt signaling activity in a specific cell type (18). Within the kidney cortex, we observed
that the MD cells have highest nuclear GFP expression as compared to the other cell types (Fig.
3.1A). Moreover, immunolabeling for active β-catenin (ABC) in wildtype (WT) mouse kidney
tissue sections also showed high expression across the entire MD cell plaque, within both the
nuclear and cytoplasmic compartments (Fig.3.1B). Finally, in the human MD,
immunohistochemistry data from the Human Protein Atlas (HPA) database (21) demonstrated the
MD-specific enrichment of Wnt pathway proteins including glycogen synthase kinase 3β
(GSK3β), TCF4, and LEF1. Taken together, these data from both the mouse and human kidney
tissue highlighted the robust level of Wnt signaling activity within the MD cell plaque.
Next, we used a neuronal nitric oxide synthase (nNOS)/Cre based approach to alter Wnt
signaling activity in an inducible fashion specifically in the MD cells (2). Since β-catenin is the
downstream effector of the canonical Wnt pathway (9), we utilized 2 genetic mouse strains in
which β-catenin is floxed at specific positions. In the gain-of-function (gof) mouse model-MD-
Wnt
gof
- the excision of exon 3 upon tamoxifen induction results in the MD-specific expression of
the stabilized form of β-catenin; exon 3 encodes the phosphorylation site target of GSK3β (16).
Alternatively, the excision of exons 2-6 of β-catenin in the MD-Wnt
lof
mice (17) results in the
deletion of β-catenin in the MD cells upon induction. We then functionally validated these mouse
strains by backcrossing them with the WntGFP reporter mouse (18). Endogenous WntGFP
81

expression in the kidney tissue of MD-Wnt
gof
:WntGFP mice showed an increase in GFP intensity
specifically in the MD plaque, demonstrating increased Wnt signaling activity as compared to
control mice (Fig.3.2D). Consistent with the deletion of β-catenin in the MD cells, in the MD-
Wnt
lof
:WntGFP mice, GFP expression was abolished specifically in the MD cell plaque (Fig.3.2C).
This methodology helped to validate the specificity of Cre-mediated excision and manipulation of
Wnt signaling activity in MD cells. The development of these MD-specific mouse models
provided us valuable tools to modulate Wnt signaling activity in MD cells.
Detailed morphological phenotyping of the kidney tissue of both the MD-Wnt
gof
and MD-
Wnt
lof
mice showed robust alterations in glomerular morphology compared to that in control WT
mice. Upregulation of MD cell Wnt signaling led to the increased size and podocyte and
endothelial cell densities of glomeruli, while hypomorphic glomeruli were observed in mice with
diminished MD cell Wnt signaling (Figs. 3.3 and 3.6). These results confirmed the essential and
non-traditional role of MD Wnt signaling activity driving vascular and glomerular endogenous
tissue remodeling, consistent with a newly discovered master regulatory role of MD cells
functioning as the nephron central command. In addition, these glomerular tissue morphological
alterations were consistent with the observed changes in glomerular function as discussed below.
As a part of the juxtaglomerular apparatus (JGA), the critical role of MD cells in regulating
GFR, blood pressure (BP), and renin expression has been established several decades ago (3, 6).
Keeping with this classic role of MD cells in the kidney, we investigated the effect of the MD Wnt
pathway on global GFR using the MediBeacon transdermal sensor technique. Upregulated MD
Wnt signaling significantly increased GFR in the MD-Wnt
gof
mice compared to control mice while
GFR was significantly lower in response to downregulated MD Wnt signaling (Fig. 3.3H). To our
knowledge, these experiments provided for the first time evidence for the regulation of GFR by
82

the MD Wnt signaling pathway in addition to the traditional role of the mitogen activated protein
kinase (MAPK) pathway in these cells (6). Thus, alterations in MD Wnt signaling have both
structural and functional consequences in the kidney.
Renin synthesis and secretion from the juxtaglomerular (JG) cells is primarily regulated by
autocoids-prostaglandin E2 (PGE2) and nitric oxide (NO)- synthesized by MD cells (1).
Immunolabeling for renin at the glomerular vascular pole showed an almost 100% increase in the
number of renin
+
cells in the MD-Wnt
gof
mice (Fig. 3.4G). Moreover, immunoblots to ascertain
total renin expression in the kidney cortex showed that MD cell Wnt signaling promotes renin
expression in the kidney cortex. Since we did not observe any change in the MAPK pathway, these
experiments highlighted the importance of the Wnt pathway in the direct regulation of well-
established MD cell functions.
As mentioned earlier, the Wnt pathway can regulate protein expression via the mTOR
signaling pathway (12). We probed the autocrine effect of Wnt signaling on MD protein synthesis
using the previously established OPP labeling technique (15, 22) and as described in Chapter 2. In
MD-Wnt
gof
mice, quantification of OPP intensity revealed approximately a 40% increase in overall
rate of protein synthesis (Fig.3.5E); however, in the MD-Wnt
lof
mice, no change was observed as
compared to control mice. Since the central regulator of cellular protein synthesis is the mTOR
protein complex (27), unchanged overall protein synthesis activity in the MD-Wnt
lof
mice can be
attributed to the normally low activity of the mTOR complex. Experiments from our laboratory
(described in Chapter 2) have demonstrated the elevated basal protein synthesis activity in MD
cells, driven by mTOR-mediated protein translation. Additionally, physiological activation of MD
cells using low salt (LS) diet increased protein synthesis activity across all mouse strains; this
effect was particularly significant in case of the MD-Wnt
gof
mice (Fig. 3.5G).
83

To determine specific proteins whose expression could be altered via Wnt signaling, we
analyzed the MD cell gene profile recently established by our laboratory (unpublished data) and
identified several angiogenic and glomerulotrophic proteins and peptides that are regulated by the
Wnt pathway. One such protein that has a potent effect on endothelial cells is CCN family member
1 (CCN1) (23, 24). As a proof-of-concept, we measured CCN1 expression in kidney cortex
homogenates of MD-Wnt
gof
and MD-Wnt
lof
mice. CCN1 expression was significantly lower in
MD-Wnt
lof
mice which is consistent with studies that have demonstrated Wnt-regulation of CCN1
expression (Fig. 3.5H) (23). While MD-Wnt
gof
mice tended to have higher CCN1 expression as
compared to control mice, it was not statistically significant, potentially due to the high Wnt
signaling activity in MD cells even in WT mice. In addition to CCN1, MD cell gene profile also
showed that several other angiogenic and tissue remodeling peptides are significantly enriched in
MD cells (unpublished data). The presence of secretory organelles at the basal surface of MD cells
(3) combined with this high level of enrichment of angiogenic and tissue remodeling peptides
suggests that in addition to the traditional function of MD cells in the kidney, these cells could
play a novel role in mediating local tissue remodeling. Several data from our laboratory using
novel progenitor cell mouse models have underscored the role of MD cells in recruiting resident
progenitor cell populations (unpublished data). To substantiate this hypothesis, we quantified
changes in the number of podocytes and glomerular endothelial cell populations in 3D cleared
kidney tissue. We observed that MD Wnt signaling activity has a potent effect on both these cell
populations; podocyte and endothelial cell number was higher in the glomeruli of MD-Wnt
gof
mice
while the opposite effect was observed in MD-Wnt
lof
mice (Fig. 3.6). These observations are
consistent with the newly identified non-traditional role of MD cells in regulating glomerular
remodeling.
84

In summary, the MD cell Wnt/β-catenin signaling pathway plays a critical role in mediating
the traditional JGA and non-traditional structural and functional tissue remodeling functions of
MD cells in the kidney. Further studies are needed regarding the regulation, secretion, and
mechanism of action of MD-specific tissue remodeling factors and their potential role in
endogenous tissue repair to fully understand this novel aspect of MD cell biology.

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Chapter 4

Disease modifying role of macula densa cell Wnt signaling activity in diabetic
kidney injury

4.1. Abstract
Although diabetic nephropathy (DN) is the leading cause of end stage renal disease
(ESRD), specific therapies are lacking due to the incomplete understanding of disease
pathogenesis. DN development and progression are characterized by initial hypertrophy,
hyperfiltration, and renin-angiotensin system (RAS) activation followed by albuminuria
development, declining glomerular filtration rate (GFR), glomerulosclerosis, and eventually
nephron loss. Within the nephron, the macula densa (MD) regulates renal hemodynamics, GFR,
and renin expression in addition to sensing local tissue metabolites. Recent clinical results with
sodium-glucose cotransporter 2 (SGLT2) inhibitors pointed to the key role of MD cells in DN
pathogenesis. In addition, our new work demonstrated that the MD Wnt signaling pathway has a
robust effect on glomerular structure, renal function, and renin expression (Chapter 3). The present
study tested the hypothesis that MD Wnt signaling activity has disease modifying roles in DN by
using the streptozotocin (STZ)-induced Type I diabetes mouse model and novel transgenic mouse
strains with MD-specific Wnt gain-of-function (gof) (MD-Wnt
gof
) and loss-of-function (lof) (MD-
Wnt
lof
). The low dose STZ regimen successfully resulted in the induction of hyperglycemia in all
mouse strains. STZ-treated mice showed a significant increase in total kidney weight, with the
effect being most pronounced in the MD-Wnt
gof
mice. Massons’ Trichrome staining-based tissue
fibrosis quantification showed a dramatic increase in fibrosis along with a progressive increase in
88

albuminuria in response to upregulated MD Wnt signaling and hyperglycemia. In contrast, the
development and progression of DN was significantly attenuated in MD-Wnt
lof
mice. The present
study uncovered the important role of the MD Wnt signaling pathway in a pathophysiological
context, providing a potential target to develop specific, mechanism-based, and highly efficient
therapies to treat DN.
Keywords: macula densa, diabetic nephropathy, streptozotocin, albuminuria, fibrosis

4.2 Introduction
Diabetic kidney disease (DKD) is the leading cause of end-stage renal disease (ESRD) in
the United States, affecting nearly 30% of patients with Type 1 diabetes mellitus (DM) and 40%
of patients with Type 2 DM (1-3). The trajectory of DKD progression is characterized by
glomerular hyperfiltration and hypertrophy followed by a progressive increase in albuminuria,
glomerulosclerosis, reduction in glomerular filtration rate (GFR), increased cardiovascular risk,
and finally, ESRD. Structural changes typical of DKD include a thickening of the glomerular
basement membrane (GBM), mesangial matrix expansion, progressive loss of podocytes, and
ultimately glomerulosclerosis and tubulointerstitial fibrosis. Risk factors for the development of
DKD are varied and include hyperglycemia, hypertension, obesity, and certain dietary factors (4).
Since DKD is a progressive disease, most people with DKD will ultimately require renal
replacement, unless new and highly efficient therapies are developed that can stop or reverse
kidney function decline. Presently, only non-specific therapies are available, including the current
standard of care, renin-angiotensin system (RAS) blockers. Even the newest drugs from recent
successful clinical trials, sodium-glucose cotransporter 2 (SGLT2) inhibitors, and glucagon-like
89

peptide 1 (GLP 1) agonist can only slow down chronic kidney disease (CKD) progression to ESRD
(5). Clearly, there continues to be an urgent and unmet medical need for specific and highly
efficient CKD therapies for treatment of millions of patients worldwide. Their development
depends on clear mechanistic understanding of DKD pathogenesis.
Persistent hyperglycemia associated with DKD results in massive reuptake of sodium and
glucose in the proximal tubule (PT), thereby dramatically reducing the amount of salt delivered to
the macula densa (MD). This in turn results in the inhibition of the tubuloglomerular feedback
(TGF), resulting in dilatation of the afferent arteriole (AA), increased glomerular capillary (GC)
pressure, and mechanical strain on podocytes, and glomerular hyperfiltration (6). Damage to the
GBM and podocytes due to the increased filtration pressure eventually results in podocyte and
nephron loss. The use of the recently developed SGLT2 inhibitors prevents sodium and glucose
reuptake in the PT, thereby increasing salt delivery to the MD and restoring TGF, and reducing
glomerular mechanical strain. DKD is also characterized by continuous activation of the RAS;
however, the renoprotection offered by RAS inhibitors in the context of nondiabetic kidney disease
is not as effective in the case of DKD, mainly due to TGF inhibition seen in DKD (6). Previous
studies from our laboratory and others have also described the diabetes relevance of several MD
cell molecular mechanisms including nitric oxide signaling, sodium-glucose cotransporter 1
(SGLT1), and the presence of the G-protein coupled receptor 91 (GPR91) on the luminal surface
of MD cells that is activated upon the binding of its ligand succinate (7-11). Using a Type I DM
mouse model, our laboratory has shown GPR91-dependent activation of the classic MD mitogen
activated protein kinases (MAPKs)-p38 and extracellular-signal related kinase 1/2 (ERK1/2) along
with increased renin release from the juxtaglomerular (JG) cells. GPR91 deletion inhibited this
renin release, suggesting that this mechanism could play an integral role in the activation of RAS
90

in DKD (12). In addition, a classic and commonly recognized renal histopathological finding in
DKD patients is the ongoing neo-angiogenesis at the glomerular vascular pole that leads to the
formation of numerous aberrant small arterioles and capillary vessels at the base of MD cells (13).
In light of the newly discovered high-rate synthesis of angiogenic proteins in MD cells (as
described in Chapters 2 and 3) it is likely that MD cells directly and primarily contribute to this
classic pathological feature of DKD.
The development of SGLT2 inhibitors, the presence of metabolite receptors in MD cells
and persistent RAS activation in DKD has underscored the important role of the MD and TGF in
mediating the progression or regression of DKD (6). However, the exact mechanisms and roles of
MD cells in the diabetic milieu are incompletely understood. Recent work from our laboratory has
shown that the MD Wnt/β-catenin signaling pathway plays an important regulatory role with
respect to the JGA functions of MD cells along with promoting glomerular tissue remodeling in
the healthy kidney (unpublished data, Chapter 3). The present study tests the hypothesis that the
MD Wnt signaling pathway has a central, disease modifying role in the context of the development
and progression of DKD pathophysiology via its major effects on the structural and functional
integrity of the glomerular filtration barrier.

4.3 Materials and Methods
4.3.1 Animals
All animal protocols were approved by the Institutional Animal Care and Use Committee
at the University of Southern California. Tamoxifen-inducible, conditional MD-Wnt
gof,
and MD-
Wnt
lof
mice were generated using nNOS/CreERT2 mice (Jackson Laboratory, Bar Harbor, ME)
for MD-specificity (14). MD-Wnt
gof
mice on a mixed background were generated by intercrossing
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nNOS/CreERT2 mice with β-catenin/fl mice (exon 3 of β-catenin flanked by loxP sites) (Jackson
Laboratory) (15) while MD-Wnt
lof
mice on a C57BL6/J background were generated by
intercrossing nNOS/CreERT2 mice with β-catenin/fl mice (exons 2-6 of β-catenin flanked by loxP
sites) (Jackson Laboratory) (16). Both strains were further backcrossed with mTmG/fl reporter
mice (Jackson Laboratory) for expression of the membrane targeted dual color fluorescent Cre-
reporter tandem dimer Tomato (mT, tdTomato) and enhanced green fluorescent protein (mG,
eGFP). Tamoxifen induction results in MD-specific alterations in β-catenin expression along with
expression of eGFP. Tamoxifen-inducible, conditional control MD-GFP mice on a C57BL6/J
background were generated as described previously (14) by intercrossing nNOS/CreERT2 and
mTmG/fl mice (Jackson Laboratory).
4.3.2 Treatments
Mice were induced using 75 mg/kg body weight tamoxifen (Alfa Aesar, Haverhill, MA)
via oral gavage for a total of 3 times every alternate day. Male mice were induced at 6 weeks of
age followed by 2 weeks of tamoxifen washout. Low dose streptozotocin (STZ) (MilliporeSigma,
Burlington, MA) treatment was carried out following Animal models of diabetes complications
consortium (AMDCC) protocols as described previously (17). Briefly, 8 week old mice were
intraperitoneally injected with 50 mg/kg body weight STZ dissolved in 0.05M sodium citrate
buffer (pH 4.5) for 5 consecutive days, and tissue was harvested 8 weeks post induction of
hyperglycemia.
4.3.3 Tissue processing and histology
Mouse kidney tissue was harvested after anesthetizing mice with 100 mg/kg body weight
ketamine and 10 mg/kg body weight xylazine and perfusing ice-cold 1X phosphate-buffered saline
(PBS) and 4% paraformaldehyde (PFA) for 2 minutes using a peristaltic pump. Tissue was fixed
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using 4% PFA for 2 hours at room temperature followed by dehydration overnight at 4℃ in 70%
ethanol. Tissue was embedded in paraffin wax and sectioned 8µm thick. Masson’s Trichrome stain
kit (Polysciences, Inc., Warrington, PA) for Trichrome staining of kidney tissue sections and
quantified by assigning Trichrome scores (18). Tissue sections were imaged at 20X magnification
using bright field microscopy.
4.3.4 Measurement of albuminuria and blood glucose
Blood glucose levels were measured post STZ treatment using Contour glucose test strips
(Bayer AG, Leverkusen, Germany) to confirm induction of hyperglycemia; blood glucose > 250
mg/dL was considered as hyperglycemia. Follow-up measurements were done after 4 weeks of
hyperglycemia and before tissue harvest. Urine was collected at baseline, after 4 weeks of
hyperglycemia, and before tissue harvest. Urine albumin was measured using Exocell Albuwell M
Mouse Albumin ELISA kit and urine creatinine was measured using Exocell Creatinine
Companion (Ethos Biosciences, Inc., Newtown Square, PA).

4.4 Results
4.4.1 Generation of diabetic kidney disease in control WT, MD-Wnt
lof
, and MD-Wnt
gof

mice
Using the low dose streptozotocin (STZ) treatment, we first aimed to establish Type I
diabetes mellitus (DM) to study the early stages of diabetic kidney disease (DKD) (17). The low
dose STZ regimen is a simple and reproducible method that can be used to model the change that
occur in the earliest stages of DKD. Control, MD-Wnt
lof
, and MD-Wnt
gof
mice were first induced
using tamoxifen at 6 weeks of age and then were either treated with the STZ drug or the vehicle
control for 5 consecutive days. We then confirmed successful induction of hyperglycemia by
93

measuring blood glucose in these mice; blood glucose > 250 mg/dL was considered as
hyperglycemic. As shown in Fig. 4.1, the low dose STZ regimen resulted in a significant elevation
of blood glucose as compared to baseline in all groups of mice. Blood glucose levels post STZ
treatment on average were as follows: control (443 ± 41 mg/dL, n = 7, p < 0.0001), MD-Wnt
lof

(464 ± 18 mg/dL, n = 17, p < 0.0001) and MD-Wnt
gof
(510 ± 20 mg/dL, n = 12, p < 0.0001) mice.
No change was observed in mice that were treated with vehicle: control (180 ± 12 mg/dL, n = 10),
MD-Wnt
lof
(179 ± 13 mg/dL, n = 11) and MD-Wnt
gof
(144 ± 6 mg/dL, n = 5).

4.4.2 Phenotypic and histological characterization of DKD mouse models
We first measured changes in the total kidney weight of mice treated either with vehicle or
STZ. As quantified in Fig. 4.2, we observed an increase in the normalized kidney weight in mice
that were treated with STZ 8 weeks post hyperglycemia (control: 0.01576 ± 0.0010, n = 9; MD-
Wnt
lof
: 0.01612 ± 0.0005, n = 16; MD-Wnt
gof
: 0.01924 ± 0.0011, n = 8) compared to the respective
Figure 4.1. Measurement of blood glucose levels in healthy and diabetic wildtype control, MD-Wnt
lof
, and MD-Wnt
gof
mice. Statistical summary of blood glucose measurements of control (n = 7-10), MD-Wnt
lof
(n = 11-17), and MD-Wnt
gof
mice
(n = 5-12) treated with vehicle (Veh) or streptozotocin (STZ) at baseline and at study endpoint. Data are expressed as mean
± SEM, ****p< 0.0001, two-way ANOVA with Šidák’s multiple comparisons test.
94

mice treated with vehicle (control: 0.01328 ± 0.0003, n = 15; MD-Wnt
lof
: 0.01294 ± 0.0004, n =
11, p = 0.0022; MD-Wnt
gof
: 0.01141 ± 0.0007, n = 5, p < 0.0001). Moreover, this effect was
markedly higher in the glycemic MD-Wnt
gof
mice (0.01924 ± 0.0011, n = 8) as compared to the
glycemic control mice (0.01576 ± 0.0010, n = 9, p = 0.0091).
Additionally, to visualize and quantify changes in the renal structure in response to 8 weeks
of hyperglycemia, we stained kidney tissue sections with Masson’s Trichrome stain to observe
glomerulosclerosis and tissue fibrosis. All kidney tissue sections were stained simultaneously and
then were assigned a fibrosis score in a blinded fashion as described before (18). Multiple (4-5)
regions of interest (ROIs) were imaged per sample. Trichrome staining of kidney tissue sections
showed the presence of glomerular mesangial expansion and interstitial fibrosis in STZ vs control
groups as shown in representative images (Fig. 4.3A-F). Interestingly, unlike in other experimental
Figure 4.2. Effects of genetic manipulations of MD Wnt signaling on kidney weight. Statistical summary of kidney weight
normalized to body weight of control (n = 9-15), MD-Wnt
lof
(n = 11-16), and MD-Wnt
gof
(n = 5-8) mice treated with vehicle
(Veh) or streptozotocin (STZ) after 8 weeks of glycemia. Data are expressed as mean ± SEM, *p<0.05, **p<0.01, ****p<
0.0001, one-way ANOVA with Šidák’s multiple comparisons test.
95

groups, nodular-like segmental glomerulosclerosis developed in STZ-treated MD-Wnt
gof
mice
(Fig. 4.3F).
Treatment with STZ resulted in a significant elevation in tissue fibrosis across all groups
of mice (control: 2.5 ± 0.2, n = 4; MD-Wnt
lof
: 2.1 ± 0.1, n = 4; MD-Wnt
gof
: 3.8 ± 0.2, n = 4) as
compared to the respective mice treated with vehicle (control: 1.2 ± 0.1, n = 4, p < 0.0001; MD-
Wnt
lof
:

1.4 ± 0.1, n = 4, , p = 0.0015; MD-Wnt
gof
: 2.3 ± 0.1, n = 4, , p < 0.0001). However, like the
Figure 4.3. Histological characterization of kidney tissue in healthy and diabetic wildtype control, MD-Wnt
lof
, and
MD-Wnt
gof
mice. A-F. Representative images of mouse kidney tissue sections of control (A-B), MD-Wnt
lof
(C-D), and
MD-Wnt
gof
mice (E-F) treated with vehicle (Veh) or streptozotocin (STZ) (n = 4) with Masson’s Trichrome staining after
8 weeks of glycemia. G. Statistical summary of Trichrome staining-based histological scoring of kidney tissue fibrosis and
glomerulosclerosis after 8 weeks of glycemia. In each tissue section, 4-5 regions of interest (ROIs) were placed and scores
were averaged Data are expressed as mean ± SEM, **p<0.01, ****p< 0.0001, one-way ANOVA with Tukey’s multiple
comparisons test.
96

trend observed with respect to changes in kidney weight, fibrosis scores were significantly higher
in glycemic MD-Wnt
gof
mice (3.8 ± 0.2, n = 4) as compared to glycemic control mice (2.5 ± 0.2, n
= 4, p <0.0001).
4.4.3 Changes in albuminuria due to induction of DKD
In order to quantify changes in albuminuria due to STZ-induced DKD, we measured
changes in the urine albumin-to-creatinine ratio (ACR) at baseline, 4 weeks post hyperglycemia,
and before tissue harvest. As quantified in Fig. 4.4, at baseline there was no significant difference
in the urine ACR across all mouse strains. 4 weeks of hyperglycemia significantly elevated urine
ACR in the STZ treated MD-Wnt
gof
mice (1861 ± 654 µg albumin/mg creatinine, n = 6) as
compared to vehicle treated MD-Wnt
gof
mice (112 ± 48 µg albumin/mg creatinine, n = 7, p =
0.0065). Interestingly this increase in ACR in glycemic MD-Wnt
gof
mice (1861 ± 654 µg
albumin/mg creatinine, n = 6) was significantly higher even compared to STZ treated control mice
(303 ± 142 µg albumin/mg creatinine, n = 6, p = 0.0139). Additionally, after 8 weeks of
hyperglycemia, this effect persisted in the glycemic MD-Wnt
gof
mice (1940 ± 941 µg albumin/mg
creatinine, n = 10) as compared to glycemic control mice (357 ± 221 µg albumin/mg creatinine, n
= 9, p = 0.0005). No change was observed in MD-Wnt
lof
mice after STZ treatment after either 4
weeks or 8 weeks.
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4.5 Discussion
The present study used a streptozotocin (STZ)-induced Type I diabetes mellitus (DM)
mouse model to study alterations in the development and initial stages of diabetic kidney disease
(DKD) in the context of genetic manipulations in macula densa (MD) Wnt/β-catenin signaling
activity. Using a well-established low dose STZ regimen (17), we were able to successfully induce
hyperglycemia in control, MD-Wnt
gof,
and MD-Wnt
lof
mice. Tissue phenotyping was carried out
using Masson’s Trichrome staining which revealed significant increase in kidney tissue fibrosis in
response to hyperglycemia. This effect was most potent in the STZ treated MD-Wnt
gof
mice as
Figure 4.4. Measurement of albuminuria in response to altered MD Wnt signaling. Statistical summary of urine
albumin-to-creatinine ratio (ACR) of control (n = 6-13), MD-Wnt
lof
(n = 10-14), and MD-Wnt
gof
(n = 4-10) mice treated
with vehicle (Veh) or streptozotocin (STZ) at baseline, and after 4 and 8 weeks of glycemia. Data are expressed as mean
± SEM, *p<0.05, **p<0.01, ***p<0.001, ****p< 0.0001, two-way ANOVA with Tukey’s multiple comparisons test.
98

compared to control and MD-Wnt
lof
mice. Additionally, albuminuria also trended similar to tissue
fibrosis, with MD-Wnt
gof
mice displaying a significant increase in albuminuria after establishment
of hyperglycemia which was not observed in the other mouse strains. Taken together, this study
provides a new insight regarding the potential involvement and importance of the MD Wnt
signaling pathway in the development and progression of DKD.
DM is a highly complex disease affecting multiple organ systems including the
cardiovascular and renal systems. However, the kidney is the primary site of DM induced
microvascular damage (4). First identified by Mogensen due to the presence of small quantities of
albumin in the urine (19), the progression of DKD is not uniform amongst Type I or Type II DM
patients. The pathophysiology of DKD is mainly driven by the hyperglycemia-induced cellular
dysfunction in the kidney and is characterized by glomerulosclerosis, glomerular hypertrophy,
tubulointerstitial fibrosis, and eventually nephron loss (20). The involvement of multiple factors
in the development of pathophysiology makes it especially complicated to identify relevant targets
to develop treatments and therapies. The early stages of DKD are characterized by glomerular
hyperfiltration, impaired tubuloglomerular feedback (TGF), and persistent activation of the renin-
angiotensin system (RAS) (6). A key cell type that plays a regulatory role in each of these processes
is the MD cell (6, 8, 10). Thus, the focus of the present study was to discern a potential mechanism
that could drive the overall involvement of MD cells in the context of DKD.
Previous studies from our laboratory have demonstrated the major role of the Wnt/β-
catenin signaling in promoting renin release from juxtaglomerular (JG) cells, elevating glomerular
filtration rate (GFR), and most importantly, mediating glomerular tissue remodeling. Our studies
also showed that the MD Wnt signaling pathway promotes overall rate of protein synthesis in the
MD cells including the synthesis of specific angiogenic and tissue remodeling peptides at the basal
99

surface (Chapter 3). This suggested that the pathological and persistent activation of this signaling
pathway in MD cells could potentially exacerbate a disease condition and vice versa. To explore
this hypothesis, we first established hyperglycemia in control, MD-Wnt
gof
, and MD-Wnt
lof
mice
using STZ. Although STZ-induced DKD does not manifest the entire pathophysiology of DKD, it
is a useful model system to study the initial stages of DKD. Moreover, the use of a low dose STZ
regimen prevents the non-specific toxic effects of STZ. STZ treatment causes cytotoxic damage
to the pancreatic β cells, resulting in the development of hyperglycemia (17). In all our mouse
strains, 5 days of STZ treatment resulted in significant elevation of blood glucose (approximately
450 mg/dL) as compared to mice treated with the vehicle. Hyperglycemia persisted over the course
of 8 weeks of follow-up.
In addition to the measurement of blood glucose, assessment of progression of DKD can
be monitored by measuring several factors- GFR, albuminuria, blood pressure (BP), and
histopathological analysis of kidney tissue. In the present study, we used the Masson’s Trichrome
stain to visualize collagen fibrils and score tissue fibrosis (18). Tissue fibrosis scores varied
significantly across the 3 mouse strains. Hyperglycemia significantly increased tissue fibrosis
across all 3 mouse strains; however, this effect was exacerbated in the MD-Wnt
gof
mice. These
results suggest that the upregulation of Wnt signaling specifically in MD cells exacerbates the
development and progression of tissue fibrosis in DKD. Conversely, inducible downregulation of
Wnt signaling in adult MD-Wnt
lof
mice reduced DKD pathology including protection of
albuminuria (Fig. 4.4) and glomerulosclerosis development (Fig. 4.3C, D, G). The hypomorphic
glomeruli with reduced cellularity observed in MD-Wnt
lof
mice as shown here and in our previous
study (unpublished data, Chapter 3) are consistent with the previously reported findings with renal
epithelial cell-specific β-Catenin/Wnt signaling deficiency (21). Pax8-driven β-catenin deficiency
100

resulted in hypoplastic renal cortex, and small under-developed glomeruli with the appearance of
parietal podocytes based on genetic lineage switch between parietal epithelial cells and podocytes
(21). Interestingly, MD-Wnt
lof
mice also feature parietal podocytes based on WT-1 labeling (Fig.
3.6A) and all of the above glomerular morphological alterations. These results suggest that Wnt
signaling in MD cells rather than in parietal epithelial cells were responsible for the previously
observed glomerular pathologies and further underscore the novel and central role of MD cells in
glomerular tissue remodeling. However, further characterization of these findings using other
histological stains like periodic acid-Schiff (PAS) staining to assess mesangial expansion and
transmission electron microscopy to assess thickening of glomerular basement membrane (GBM)
is required to completely describe the effect of upregulated MD Wnt signaling.
Another important indicator of DKD is the progressive increase in albuminuria (17, 20).
Using ELISA kits to measure urine albumin and creatinine, we quantified changes in the urine
albumin-to-creatinine ratio (ACR) at baseline, 4 weeks after hyperglycemia, and before tissue
harvest. As expected, urine ACR was unchanged at baseline; however, 4 weeks after inducing
hyperglycemia, ACR was significantly higher specifically in the STZ treated MD-Wnt
gof
mice
(Fig. 4.4). After 8 weeks of hyperglycemia, this effect persisted in the MD-Wnt
gof
mice while STZ
treated control mice, there was slight but non-significant increase in albuminuria. No change was
observed in the MD-Wnt
lof
mice which had the lowest ACR compared to the other mouse strains
(Fig. 4.4). These results are relevant to the classic clinical phenotype of DKD patients
(albuminuria) and entirely consistent with the observed histopathological alterations (Fig. 4.3)
suggesting the disruption of the glomerular filtration barrier in STZ-induced hyperglycemia and
most importantly, its exacerbation in MD-Wnt
gof
mice.
101

The current study provided a preliminary analysis of specific aspects of the role of the MD
Wnt signaling pathway in a pathophysiological context. Using MD-specific inducible genetic
mouse models, we provided the first evidence that upregulated MD Wnt signaling exacerbates the
development of DKD, while MD cell Wnt inhibition has potential protective effects in DKD.
Further morphological and functional phenotyping combined with analysis of the molecular
mechanism responsible for this effect is required for detailed assessment of this phenomenon. Such
a complete analysis would provide key evidence regarding the role of MD cells in the initial stages
of DKD and potentially lead to the development of targeted, more efficient therapies to treat DKD.

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2019.

12. Vargas SL, Toma I, Kang JJ, Meer EJ, and Peti-Peterdi J. Activation of the succinate
receptor GPR91 in macula densa cells causes renin release. J Am Soc Nephrol 20(5): 1002-
11, 2009.

13. Löwen J, Gröne E, Gröne HJ, and Kriz W. Herniation of the tuft with outgrowth of vessels
through the glomerular entrance in diabetic nephropathy damages the juxtaglomerular
apparatus. Am J Physiol Renal Physiol 317(2): F399-410, 2019.

14. Gyarmati G, Shroff UN, Riquier-Brison A, Kriz W, Kaissling B, Neal CR, Arkill KP,
Ahmadi N, Gill IS, Moon JY, Desposito D, and Peti-Peterdi J. A new view of macula
densa cell microanatomy. Am J Physiol Renal Physiol 320(3): F492-504, 2021.

15. Harada N, Tamai Y, Ishikawa T, Sauer B, Takaku K, Oshima M, and Taketo MM.
Intestinal polyposis in mice with a dominant stable mutation of the beta-catenin gene.
EMBO J 18(21): 5931-42, 1999.

16. Brault V, Moore R, Kutsch S, Ishibashi M, Rowitch DH, McMahon AP, Sommer L,
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deletion results in dramatic brain malformation and failure of craniofacial development.
Development 128(8): 1253-64, 2001.

103

17. Breyer MD, Böttinger E, Brosius FC 3
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2005.

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104

Chapter 5

Conclusions and future directions

5.1 Summary of dissertation project
In this dissertation project, we discussed several newly discovered cell biological and
physiological mechanistic features of the mysterious but chief renal cell type of the macula densa
(MD), and their roles in regulating key renal and glomerular functions in health and disease.
Specifically, the main focus was the newly uncovered high activity and new roles of 2 major
Figure 5.1. Summary of the current dissertation project.
Created with Biorender.com
105

signaling pathways- the mammalian target of rapamycin (mTOR) and Wnt/β-catenin signaling
pathways- in MD cells and their downstream autocrine and paracrine effects on renal structure and
function (Fig. 5.1).
In Chapters 2 and 3, using MD-specific inducible genetic mouse models and novel cell
biological tools, we established the importance of both the mTOR and Wnt/β-catenin signaling
pathways regulating the protein synthesis capacity of MD cells. These experiments clearly
demonstrated the high level of protein synthesis within the MD cells under control conditions.
Moreover, upregulation of both the MD mTOR and Wnt signaling pathways resulted in a
significant increase in global MD protein synthesis as well in the expression of several MD-
enriched tissue remodeling and angiogenic peptides (Pappa2, CCN1, CxCl14, etc.)
Figure 5.2. Schematic depicting newly identified relevant MD cell signaling pathways.

Created with Biorender.com
106

The use of intravital multiphoton microscopy (MPM) and other imaging approaches
provided an unprecedented and detailed view of the glomerular vascular pole region in these new
mouse models and allowed us to quantitatively visualize the changes in renal and glomerular
hemodynamics in response to elevated MD protein synthesis in the intact, living kidney.
Upregulated MD mTOR signaling resulted in elevated glomerular filtration rate (GFR) and renal
blood flow (RBF) within the glomerular capillaries (GCs) as well as the afferent (AA) and efferent
(EA) arterioles. Altered MD Wnt signaling activity also affected GFR; increased MD Wnt
signaling elevated GFR with the opposite effect in response to downregulated MD Wnt signaling.
Using these newly established mouse models, we also addressed the effect of these
signaling pathways in regulating the classic, traditional function of MD cells within the
juxtaglomerular apparatus (JGA), namely the regulation of renin expression and renal and
glomerular hemodynamics. Increased MD mTOR and Wnt signaling dramatically elevated renin
expression along with an increase in classic MD-specific enzymes like cyclooxygenase 2 (COX2).
However, in both cases, we did not detect any changes in the mitogen activated protein kinases
(MAPKs) suggesting that these signaling pathways operate downstream of the MAPK pathway in
the MD cells. Finally, the current studies also uncovered for the first time, a new, non-traditional
role of MD cells, driving endogenous glomerular and vascular tissue remodeling, as a result of the
synthesis and secretion of angiogenic and tissue remodeling peptides. Altered MD Wnt signaling
resulted in observable changes in glomerular structure, especially with respect to changes in the
podocytes, endothelial cells, and endothelial progenitor cell populations (Fig. 5.2).
In Chapter 4, we discovered the effect of the MD Wnt signaling pathway in the context of
a pathophysiological setting, specifically diabetic kidney disease (DKD). Using a well-established
mouse model for DKD, we provided preliminary evidence regarding the involvement of the MD
107

Wnt signaling pathway in the development and progression of DKD. Increased MD Wnt signaling
exacerbated the progression of DKD with increases in kidney weight, tissue fibrosis as well as
albuminuria. No adverse effects were observed in response to downregulated MD Wnt signaling.
Taken together, these studies painted a new picture of MD cells functioning as the nephron
central command, and for the first time discovered their high protein synthesis capacity and its
roles in regulating glomerular hemodynamics and tissue remodeling in health and disease (Fig.
5.3).
Figure 5.3. Schematic depicting novel role of MD cells in tissue remodeling.

108

5.2 Limitations and future directions
The primary focus of the current studies was to delineate specific signaling pathways that
regulate the protein synthesis and secretion capacity of MD cells and their paracrine control of
glomerular and renal functions in health and disease. Moreover, we also aimed to provide insight
regarding the novel function of MD cells as regulators of endogenous tissue remodeling. Data from
our studies conclusively establish the high rate of protein synthesis within MD cells which have a
wide-ranging effect on specific renal cell populations as well as function. We were also able to
identify a few candidate proteins whose expression was altered in response to both the mTOR and
Wnt signaling pathways. However, further studies are required to provide more information
regarding the nature of the proteins comprising the MD secretome. Using our newly established
mouse MD cell line (unpublished data) to modulate these signaling pathway will provide a
valuable tool to identify several other proteins that are synthesized by MD cells. Moreover,
although preliminary evidence from our experiments suggest that MD cells can also secrete these
proteins from their basal surface, the exact mechanism of secretion and whether it is constitutive
or regulated is yet to be determined. Data from the MD cell gene profile (unpublished data)
suggests that MD cells express several proteins that are known elements and molecular effectors
of regulated vesicle exocytosis, for example, synaptotagmins (1). Several groups have also
reported the use of RNA interference (RNAi) to inhibit the function of a class of proteins-Rab
GTPases- that are involved in exosome biogenesis and secretion (2, 3). Thus, it would be
interesting to use these tools to carefully tease out the molecular players involved in protein
secretion and exocytosis. Additionally, after having identified the effectors of exocytosis in MD
cells, these results can be replicated in vivo using mouse models that have disrupted expression of
drivers of exocytosis.
109

In our experiments regarding the role of the mTOR pathway in the MD cells, we used
short-term systemic rapamycin (Rapa) treatment to downregulate the mTOR signaling activity.
However, to specifically downregulate mTOR signaling activity in MD cells, it is necessary to
develop an inducible genetic mouse model using a similar neuronal nitric oxide synthase
(nNOS)/Cre driven approach. Finally, since our results showed upregulation of specific
chemokines and angiogenic peptides in the newly established MD-mTOR
gof
model, this new
experimental research tool will be highly valuable in several future research projects. For example,
studies can be designed using intravital MPM, the signature research technology of our laboratory,
to quantitatively visualize and mechanistically characterize the recruitment, fate, and function of
circulating immune cell populations and resident endothelial or mesenchymal progenitor cells that
migrate towards the MD epicenter and structurally and functionally remodel the glomerulus.
Recent publications from our laboratory have clearly demonstrated the efficacy of serial MPM of
the same kidney tissue volume over several days and weeks in visualizing vascular endothelial
remodeling at the single-cell level in the healthy kidney and during conditions of glomerular injury
(4). In addition, tracking the homing and function of fluorescently labeled endogenous activated
memory T cell populations in a mouse model of lupus nephritis uncovered the essential role and
the effects of therapeutic targeting of new mechanisms in the local tissue environment that
contribute to local inflammation (5). Similar approaches could be implemented in future studies
using the MD-mTOR
gof
mice.
Finally, with respect to our experiments with the DKD mouse model, we primarily used
male mice to conduct our studies due to the well-established higher susceptibility of the male
gender to DKD in conjunction with a genetic mouse strain (C57BL6/J) that is known to be resistant
to several types of injury. However, several groups including ours have demonstrated a sexually
110

dimorphic pattern with respect to the expression of renal transporters, cell structure, and renal
function (6-8). In case of DKD, it has been established that glomerular hyperfiltration seen in the
early stages of Type I diabetes induced DKD disproportionately affects women (9) whereas, in a
mouse model of DKD, it was shown that the estrogen receptor plays a renoprotective role in the
early stages of Type I diabetes induced DKD (10). Thus, it will be essential to determine in future
studies if the phenotypes and effects that were observed in male MD-Wnt
gof
mice in the present
experiments are reproducible in female MD-Wnt
gof
mice.
In summary, this chapter highlights the key findings of the dissertation project along with
a detailed analysis of the limitations of the completed studies. It also proposes several future areas
of research that can be pursued to provide a better understanding of the role of the MD cells in the
kidney. We hope that the scientific and technical advances made as a part of this research project
can aid the development of targeted therapies for kidney disease.

5.3 References
1. Chapman ER. Synaptotagmin: A Ca
2+
sensor that triggers exocytosis? Nat Rev Mol Cell
Biol 3(7): 498-508, 2002.

2. Chiang L, Karvar S, and Hamm-Alvarez SF. Direct imaging of RAB27B-enriched
secretory vesicle biogenesis in lacrimal acinar cells reveals origins on a nascent vesicle
budding site. PLoS One 7(2): e31789, 2012.

3. Ostrowski M, Carmo NB, Krumeich S, Fanget I, Raposo G, Savina A, Moita CF, Schauer
K, Hume An, Frietas RP, Goud B, Benaroch P, Hacohen N, Fukuda M, Desnos C, Seabra
MC, Darchen F, Amigorena S, Moita LF, and Thery C. Rab27a and Rab27b control
different steps of the exosome secretion pathway. Nat Cell Biol 12(1): 19-30, 2010.

4. Desposito D, Schiessl IM, Gyarmati G, Riquier-Brison A, Izuhara AK, Kadoya H, Ber B,
Shroff UN, Hong YK, and Peti-Peterdi J. Serial intravital imaging captures dynamic and
functional endothelial remodeling with single-cell resolution. JCI Insight 6(10): 123392-
3407, 2021.
111

5. Kadoya H, Yu N, Schiessl IM, Riquier-Brison A, Gyarmati G, Desposito D, Kidokoro K,
Butler MJ, Jacob CO, and Peti-Peterdi J. Essential role and therapeutic targeting of the
glomerular endothelial glycocalyx in lupus nephritis. JCI Insight 5(19): e131252, 2020.

6. Layton AT, and Sullivan JC. Recent advances in sex differences in kidney function. Am J
Physiol Renal Physiol 316(2): F328-31, 2019.

7. Gyarmati G, Shroff UN, Riquier-Brison A, Kriz W, Kaissling B, Neal CR, Arkill KP,
Ahmadi N, Gill IS, Moon JY, Desposito D, and Peti-Peterdi J. A new view of macula densa
cell microanatomy. Am J Physiol Renal Physiol 320(3): F492-F504, 2021.
8. Veiras LC, Girardi ACC, Curry J, Pei L, Ralph DL, Tran A, Castelo-Branco RC, Pastor-
Soler N, Arranz CT, Yu ASL, and McDonough AA. Sexual dimorphic pattern of renal
transporters and electrolyte homeostasis. J Am Soc Nephrol 28(12): 3504-3517, 2017.
9. Škrtić M, Lytvyn Y, Bjornstad P, Reich HN, Scholey JW, Yip P, Sochett EB, Perkins B,
and Cherney DZI. Influence of sex on hyperfiltration in patients with uncomplicated type
1 diabetes. Am J Physiol Renal Physiol 312(4): F599-606, 2017.

10. Irsik DL, Romero-Aleshire MJ, Chavez EM, Fallet RW, Brooks HL, Carmines PK, and
Lane PH. Renoprotective impact of estrogen receptor-α and its splice variants in female
mice with type 1 diabetes. Am J Physiol Renal Physiol 315(3): F512-20, 2018.

112

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Abstract (if available)

Abstract The kidney as a whole is composed of varied cellular populations, each with its specific anatomy and function. Amongst these cell types, there exists a plaque of 20-25 specialized epithelial cells, namely, the macula densa (MD) cells at the glomerular entrance of each nephron. MD cells have long been recognized to play a critical role in regulating glomerular filtration rate (GFR), renal blood flow (RBF), and renin release via tubuloglomerular feedback (TGF) mechanisms, but the general understanding of the function of MD cells has been very limited. A careful observation of the unique microanatomy of MD cells shows the abundance of endoplasmic reticulum (ER), Golgi apparatus, and secretory granules in basal cell processes, which together suggest that MD cells are endowed with an elaborate protein synthesis machinery towards the basal surface facing the glomerulus. However, the complex three-dimensional architecture of the glomerulus and the lack of a comprehensive MD cell research toolbox have made it difficult to access and study the MD cells. The recent advancements in microscopy techniques, specifically high resolution intravital multiphoton microscopy (MPM) and the development of MD-specific transgenic mouse models now provide a new view of the mysterious MD cells and allow us to carry out a detailed analysis of the unique features and functions of MD cells in the kidney. ? The current dissertation project establishes and addresses the novel role of the high protein synthesis and secretion activity of MD cells and its glomerulotrophic effect in mediating endogenous tissue remodeling. Chapter 2 of the dissertation highlights the role of the mammalian target of rapamycin (mTOR) signaling pathway in regulating MD cell protein synthesis activity as well as its effect in regulating glomerular hemodynamics and renin expression. Chapters 3 and 4 address the effect of altered Wnt/?-catenin signaling activity in normal physiological as well as pathophysiological settings. ? In Chapter 2, we clearly demonstrate the importance of the mTOR pathway in upregulating MD protein synthesis activity using a fluorescence imaging-based approach. Upregulated MD mTOR signaling also significantly increased renin expression and classic MD signaling proteins, resulting in robust increases in both GFR and RBF. ? Chapter 3 pertains to the role of the MD Wnt signaling pathway in regulating glomerular architecture and function. Using MD-specific mouse models to either upregulate or downregulate Wnt signaling, we show several changes in the glomerular structure as well as GFR and renin expression. Moreover, we also clearly demonstrate changes in specific cell populations-podocytes and endothelial cells-which highlights the previously unknown role of MD cells in mediating glomerular tissue remodeling. ? Finally, in Chapter 4, we provide evidence for the disease modifying role of the MD Wnt signaling pathway in the development and progression of diabetic nephropathy (DN). ? In conclusion, the current dissertation provides a comprehensive overview of several newly discovered aspects of MD cell function in the kidney and its role in health and disease including in tissue regeneration and repair.

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Asset Metadata

Creator Shroff, Urvi Nikhil (author)

Core Title Novel functions of the macula densa in renal physiology and disease

School Keck School of Medicine

Degree Doctor of Philosophy

Degree Program Medical Biology

Degree Conferral Date 2021-08

Publication Date 07/24/2021

Defense Date 06/15/2021

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag glomerular filtration rate,intravital multiphoton microscopy,Kidney,macula densa,mTOR,OAI-PMH Harvest,protein synthesis,renin,tissue remodeling,Wnt

Format application/pdf (imt)

Language English

Contributor Electronically uploaded by the author (provenance)

Advisor McDonough, Alicia (committee chair), Perin-Gallandt, Laura (committee member), Peti-Peterdi, Janos (committee member)

Creator Email urvi.shroff01@gmail.com,ushroff@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-oUC15619741

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