universidade de sÃo paulo...camundongos desnutridos, indicando que o microambiente medular está...

UNIVERSIDADE DE SÃO PAULO FACULDADE DE CIÊNCIAS FARMACÊUTICAS

Programa de Pós-Graduação em Farmácia

Área de Fisiopatologia e Toxicologia

Efeitos da desnutrição proteica sobre o microambiente perivascular

medular na regulação da hematopoese

Araceli Aparecida Hastreiter

Tese para obtenção do Título de

DOUTOR

Orientador: Prof. Dr. Ricardo Ambrósio Fock

São Paulo

2019

UNIVERSIDADE DE SÃO PAULO FACULDADE DE CIÊNCIAS FARMACÊUTICAS

Programa de Pós-Graduação em Farmácia

Área de Fisiopatologia e Toxicologia




Versão Original

Tese para obtenção do Título de

DOUTOR

Orientador: Prof. Dr. Ricardo Ambrósio Fock

São Paulo

2019

Autorizo a reprodução e divulgação total ou parcial deste trabalho, por qualquer meioconvencional ou eletronico, para fins de estudo e pesquisa, desde que citada a fonte.

Ficha Catalográfica elaborada eletronicamente pelo autor, utilizando oprograma desenvolvido pela Seção Técnica de Informática do ICMC/USP e

adaptado para a Divisão de Biblioteca e Documentação do Conjunto das Químicas da USP

Bibliotecária responsável pela orientação de catalogação da publicação:Marlene Aparecida Vieira - CRB - 8/5562

H358eHastreiter, Araceli Aparecida Efeitos da desnutrição proteica sobre omicroambiente perivascular medular na regulação dahematopoese / Araceli Aparecida Hastreiter. - SãoPaulo, 2019. 148 p.

Tese (doutorado) - Faculdade de CiênciasFarmacêuticas da Universidade de São Paulo.Departamento de Análises Clínicas e Toxicológicas. Orientador: Fock, Ricardo Ambr?sio

1. Desnutrição. 2. Hematopoese. 3. Célulaendotelial. 4. Célula tronco mesenquimal. I. T. II.Fock, Ricardo Ambrósio, orientador.




Comissão Julgadora

Da

Tese para obtenção do Título de DOUTOR

Prof. Dr. Ricardo Ambrósio Fock

orientador/presidente

___________________________________________ 1o. examinador

___________________________________________ 2o. examinador

___________________________________________ 3o. examinador

São Paulo, __________ de 2019.

“Por vezes sentimos que aquilo que fazemos não é senão uma gota de água no mar.

Mas o mar seria menor se lhe faltasse uma gota”

Madre Teresa de Calcutá

DEDICATÓRIA

Aos meus pais Salete e Guido,

por todo o apoio, ensinamentos e amor incondicional.

Ao meu marido Junior,

por todo companheirismo, incentivo e carinho.

Ao meu irmão Alisson,

por acreditar no meu potencial.

Ao Prof. Dr. Ricardo Ambrósio Fock,

orientador deste trabalho,

pela confiança, ensinamentos e profissionalismo.

Minha admiração e agradecimento.

AGRADECIMENTOS

À Faculdade de Ciências Farmacêuticas da Universidade de São Paulo (FCF-USP).

À Profa. Dra. Primavera Borelli do Departamento de Análises Clínicas da FCF-USP,

por seu exemplo de profissionalismo.

Profa. Dra. Cláudia Rodrigues, pelos ensinamentos e pela oportunidade de estágio

na University of Miami.

À Maristela Tsujita e Guilherme Galvão dos Santos, pela amizade e discussões

científicas. Minha admiração e respeito pelo seu profissionalismo.

À Iara Kretzer, pela amizade indispensável.

À Renata Albuquerque e Edson Naoto Makiyama, pela amizade e auxílio técnico.

Aos companheiros de Laboratório de Hematologia Experimental, Amanda Nogueira-

Pedro, Andressa Cristina Santos, Beatriz Batista, Bruna Baptista, Carolina Carvalho

Dias, Dalila Cunha, Ed Wilson Cavalcante Santos, Graziela Batista, Jackeline Beltran,

Talita Sartori e Vaniky Duarte, pelos ensinamentos diários.

Aos funcionários da Secretaria do Departamento de Análises Clínicas da FCF-USP e

da secretaria do Programa de Pós-Graduação em Farmácia da FCF-USP, pela ajuda

prestada.

À equipe do biotério de Produção e Experimentação da FCF-USP.

À FAPESP, CAPES e CNPq pelo apoio financeiro para o desenvolvimento deste

trabalho.

Aos amigos Rodrigo Sant’Anna e Stephanie Rabbitts, por todo o apoio nessa jornada.

Aos meus lindos Leopoldo, Nicolau e Romeu, pelo amor incondicional e por me

ensinarem todos os dias que as coisas mais simples podem trazer muita felicidade.

Todo o meu amor.

VIII

RESUMO

HASTREITER, A. A. Efeitos da desnutrição proteica sobre o microambiente perivascular medular na regulação da hematopoese. 2019. 148 f. Tese (Doutorado) – Faculdade de Ciências Farmacêuticas, Universidade de São Paulo, 2019.

A desnutrição proteica (DP) provoca anemia e leucopenia decorrente da redução de precursores hematopoéticos e comprometimento da produção de mediadores indutores da hematopoese. A hematopoese ocorre na medula óssea (MO) em regiões distintas chamadas de nichos, que modulam os processos de diferenciação, proliferação e auto renovação da célula tronco hematopoiética (CTH). O microambiente perivascular, composto principalmente por células tronco mesenquimais (CTM) e células endoteliais (CE), é o principal modulador das CTH e sua função se estende até a migração das células hematopoiéticas maduras para o sangue periférico, através da produção de citocinas e fatores de crescimento. Dessa forma, nossa hipótese é que a DP altera o microambiente perivascular e objetivamos avaliar se a DP afeta a capacidade modulatória das CTM e CE sobre a hematopoese. Utilizamos camundongos C57BL/6 machos, divididos em grupos Controle e Desnutrido, sendo que o grupo Controle recebeu ração normoproteica (12% caseína) e o grupo Desnutrido recebeu ração hipoproteica (2% caseína), ambos durante 5 semanas. Após este período, os animais foram eutanasiados, foi realizada a avaliação nutricional e hematológica, caracterizando a DP. Realizamos transplantes de mielo-monoblastos leucêmicos e observamos que estas células apresentam menor taxa de proliferação e se encontram em maior quantidade nas fases G0/G1 do ciclo celular em camundongos desnutridos, indicando que o microambiente medular está comprometido. Isolamos CTM, que foram caracterizadas e diferenciadas in vitro em CE, o que foi evidenciado pelos marcadores CD31 e CD144. Quantificamos CTH e progenitores hematopoéticos, bem como reguladores de proliferação e diferenciação, ex vivo e após culturas com CTM ou CE. Observamos que a DP reduz CTH e progenitores hematopoéticos ex vivo. Na DP, as CTM promovem incremento de CTH e suprimem a diferenciação hematopoética, enquanto que as CE induzem parada no ciclo celular. Adicionalmente, observamos que a DP afeta a granulopoese por diminuição da expressão de G-CSFr nos progenitores grânulo-monocíticos. Dessa forma, concluímos que a DP compromete a hematopoese por alterações intrínsecas na CTH, como também por alterações ocasionadas no microambiente perivascular medular. Palavras-chave: Desnutrição proteica; Célula tronco mesenquimal; Célula endotelial; Regulação da hematopoese.

Palavras-chave: Desnutrição proteica; Célula tronco mesenquimal; Célula endotelial; Regulação da hematopoese.

IX

ABSTRACT

HASTREITER, ARACELI APARECIDA. Protein malnutrition effects of perivascular bone marrow microenvironment on the regulation of hematopoiesis. 2019. 148 f. Tese (Doutorado) – Faculdade de Ciências Farmacêuticas, Universidade de São Paulo, 2019.

Protein malnutrition (PM) causes anemia and leukopenia by reduction of hematopoietic precursors and impaired production of mediators that induce hematopoiesis, as well as structural and ultrastructural changes in the bone marrow (BM) extracellular matrix. Hematopoiesis occurs in the bone marrow (BM) in distinct regions called niches, which modulate the processes of differentiation, proliferation and self-renewal of the hematopoietic stem cell (HSC). The perivascular niche, composed mainly by mesenchymal stem cells (MSC) and endothelial cells (EC), is the major modulator of HSC and its function extends to the migration of mature hematopoietic cells into the peripheral blood through the production of cytokines and growth factors. Thus, our hypothesis is that PM changes the perivascular niche and our objective is to evaluate whether PM affects the modulatory capacity of MSC and EC on hematopoiesis. C57BL/6 male mice were divided into Control and Malnourished groups, which received for 5 weeks, respectively, a normal protein diet (12% casein) and a low protein diet (2% casein). After this period, animals were euthanized, nutritional and hematological evaluations were performed, featuring the PM. We performed leukemic myelo-monoblasts cells transplantation and observed that these cells have a lower proliferation rate and are rather in the cell cycle G0/G1 phases in malnourished mice, indicating that the BM microenvironment is compromised in PM. MSC were isolated, characterized and differentiated in vitro into EC cells, which were evidenced by CD31 and CD144 markers. We performed the quantification of HSC and hematopoietic progenitors, as well as some regulators of proliferation and differentiation, ex vivo and after cultures with MSC or EC. We observed that PM reduces HSC and hematopoietic progenitors ex vivo. In PM, MSC promote increase in HSC and suppress hematopoietic differentiation, whereas ECs induce cell cycle arrest. Additionally, we verified that PM affects granulopoesis by decreasing the expression of G-CSFr in granule-monocytic progenitors. Thus, we conclude that PD compromises hematopoiesis due to intrinsic alterations in HSC, as well as alterations in the medullary perivascular niche. Key-words: Protein malnutrition; Perivascular microenvironment; Mesenchymal stem cell; Endothelial cell; Hematopoiesis regulation.

X

LISTA DE ABREVIATURAS E SIGLAS

ACTH Hormônio adrenocorticotrófico AIN American Institute of Nutrition Ang Angiopoietina CAR CXCL-12 abundant reticular cell CD Cluster of differentiation CE Célula endotelial CLP Progenitor linfoide comum CMP Progenitor mieloide comum CTH Célula tronco hematopoética CTM Célula tronco mesenquimal DP Desnutrição proteica DPE Desnutrição proteico-energética EGF Fator de crescimento epidérmico FAO Food and Agriculture Organization FGF Fator de crescimento de fibroblastos Flt Receptor de VEGF tipo 1 G-CSF Fator de crescimento de colônias grânulocíticas GM-CSF Fator de crescimento de colônias grânulo-macrofágicas GMP Progenitor grânulo-monocítico HGF Fator de crescimento de hepatócitos IGF Fator de crescimento semelhante à insulina IL Interleucina Kdr Receptor de VEGF tipo 2 LepR Receptor de leptina LIN Linhagem MCAM Molécula de adesão celular de melanoma MEC Matriz extracelular MEP Progenitor megacariocítico-eritroide MPP Progenitor hematopoético multipotente MO Medula óssea OMS Organização Mundial de Saúde ONU Organização das Nações Unidas PECAM Molécula de adesão epitélio-plaquetária CLP Progenitor linfoide comum SCF Fator de células tronco SDF Fator derivado de células estromais TGF Fator de crescimento transformador TNF Fator de necrose tumoral VCAM Proteína celular de adesão vascular VEGF Fator de crescimento endotelial vascular VEGFR Receptor de VEGF vWF Fator de von Willebrand

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SUMÁRIO

1 INTRODUÇÃO ....................................................................................................... 12

1.1 DESNUTRIÇÃO .............................................................................................. 12

1.2 HEMATOPOESE E DESNUTRIÇÃO PROTEICA........................................... 14

1.3 MICROAMBIENTE MEDULAR E NICHO HEMATOPOÉTICO PERIVASCULAR .................................................................................................. 17

2 HIPÓTESE E OBJETIVOS .................................................................................... 23

3 CAPÍTULO I ........................................................................................................... 24

A DESNUTRIÇÃO PROTEICA SUPRIME A HEMATOPOESE ATRAVÉS DO COMPROMETIMENTO DAS CÉLULAS ENDOTELIAIS MEDULARES ................. 24

4 CAPÍTULO II .......................................................................................................... 62

EFEITOS DA DESNUTRIÇÃO PROTEICA SOBRE ASPECTOS REGULATÓRIOS DA HEMATOPOESE DAS CÉLULAS TRONCO MESENQUIMAIS MEDULARES 62

5 CAPÍTULO III ......................................................................................................... 93

A DIMINUIÇÃO DO RECEPTOR DE G-CSF NAS CÉLULAS PROGENITORAS GRANULOCÍTICAS CAUSA NEUTROPENIA NA DESNUTRIÇÃO PROTEICA .... 93

6 DISCUSSÃO FINAL .............................................................................................. 94

7 CONCLUSÕES .................................................................................................... 127

REFERÊNCIAS BIBLIOGRÁFICAS ...................................................................... 128

ANEXOS ................................................................................................................. 146

ANEXO I – Protocolo da Comissão de Ética no Uso de Animais .................. 146

ANEXO II – Ficha do Aluno ............................................................................... 147

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1 INTRODUÇÃO

1.1 DESNUTRIÇÃO

Segundo a Organização das Nações Unidas (ONU) e a Organização Mundial

da Saúde (OMS), todo ser humano tem direito à saúde e à nutrição adequada

(ACC/SCN, 2000), entretanto a desnutrição ainda é um dos principais problemas

alimentares no mundo.

A desnutrição é definida como uma condição fisiológica anormal, causada

pelo desequilíbrio entre a oferta e a demanda de macronutrientes (carboidratos,

proteínas e lipídeos) e/ou micronutrientes (vitaminas e minerais) essenciais para a

manutenção e desenvolvimento físico e cognitivo do organismo. Dessa forma, a

desnutrição (ou subnutrição) pode ser classificada em três categorias principais: (a)

desnutrição, (b) deficiência de micronutrientes e (c) sobrepeso e obesidade (WHO,

2011; FAO, 2012). No Brasil, o termo “desnutrição” é usualmente empregado como

sinônimo de fome ou baixo peso corporal e denota falta de um ou mais nutrientes

necessários para a saúde (STINNETT, 1983).

A desnutrição é de origem multifatorial, de forma que pode ser decorrente de

dietas baseadas em alimentos com combinações e/ou proporções nutricionais

inadequadas, bem como da deficiência de nutrientes específicos, tanto por perda ou

utilização excessiva, como ocorre em parasitoses, doenças crônicas, sepse e/ou

condições inflamatórias agudas que aumentam os processos catabólicos (SHETTY,

2003).

Dados da FAO demonstram que em 2017 havia cerca de 821 milhões de

pessoas desnutridas mundialmente, sendo que cerca de 22% das crianças abaixo de

5 anos apresentaram algum grau de desnutrição. É alarmante o fato de, após 10 anos

de declínio significativo, a incidência da desnutrição ter apresentado aumento nos

últimos três anos em mais de 37 milhões de pessoas, com maior aumento na África e

na América do Sul. Esse aumento é creditado a alterações climáticas que prejudicam

a agricultura e, principalmente, a condições socioeconômicas precárias nas regiões

de conflito e guerrilhas, que aumenta o número de pessoas em estado de extrema

pobreza (FAO, 2018).

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No Brasil, aproximadamente 5,2 milhões de pessoas apresentavam algum

quadro de desnutrição em 2017 (FAO, 2018). A prevalência da desnutrição é maior

em crianças nas regiões Norte e Nordeste (LIMA et al., 2010), mas também é

frequentemente observada nas periferias das grandes cidades de todo o território

nacional (SAWAYA et al., 2009).

A situação nutricional dos brasileiros apresentou melhora nos últimos anos,

resultante da expansão e melhorias nos serviços de saúde, saneamento básico e

programas sociais (MONTEIRO et al., 2009). Esta redução está relacionada ao

compromisso firmado do Brasil com o programa de metas dos Objetivos do Milênio da

ONU, que visa, entre outras metas, a redução da extrema pobreza e fome no mundo.

Diversos programas de combate à desnutrição, como o Programa Bolsa Família,

foram implementados ao longo dos últimos anos no Brasil. Entretanto, o estado

nutricional dos beneficiários do programa está aquém do esperado pelos objetivos do

programa, visto que quase 70% das famílias beneficiadas relataram um aumento no

consumo de alimentos altamente calóricos, mas de baixo valor nutricional (WOLF e

BARROS FILHO, 2014).

As formas mais frequentes de desnutrição advêm do déficit proteico, sendo

denominadas desnutrição proteica (DP) e proteico-energética (DPE) ou proteico-

calórica (KEUSCH, 2003a) e são definidas pela OMS como um conjunto de condições

patológicas decorrente da menor ingestão, em proporções variadas, de proteínas ou

proteínas e calorias, respectivamente (WHO, 2011). A população mais suscetível à

DP/DPE são as crianças e idosos, bem como portadores de doenças crônicas

(PEDRUZZI e TEIXEIRA, 2007).

Em pacientes hospitalizados, a desnutrição apresenta incidência de 19% a

80%, sendo que até 70% dos pacientes desnutridos no momento da admissão

hospitalar sofre uma piora gradual em seu estado nutricional, evoluindo para uma

piora do quadro clínico e, muitas vezes, comprometendo a resposta ao tratamento

(WAITZBERG, 2006; NORMAN et al., 2011). Consequentemente, a desnutrição pode

ser considerada um fator preditivo de mortalidade, como, por exemplo, para pacientes

portadores de insuficiência renal crônica sob diálise, nos quais é frequente (22 a 54%)

a instalação de DP moderada a severa (PIRATELLI e TELAROLLI JUNIOR, 2012;

VAVRUK et al., 2012).

As manifestações clínicas observadas em situações de DP ou DPE são

14

variadas e dependem da intensidade do déficit calórico e/ou proteico e sua duração,

bem como da idade do paciente, da causa da deficiência e a associação com outras

doenças (DE ANGELIS, 1986). A DP pode cursar com retardo no crescimento

(MONTEIRO et al., 2009) e provocar alterações morfológicas e funcionais nos

sistemas cardiovascular, renal, respiratório e digestório (WAITZBERG, 2006). Além

desta redução na integridade física, pode provocar efeitos psicossociais, como

depressão e ansiedade (SAUNDERS e SMITH, 2010) e diminuição de habilidades

mentais, como a função cognitiva (WHO, 2011).

Os casos extremos de DP e DPE originam duas síndromes – Marasmus e

Kwashiorkor – que podem ocorrer de forma isolada ou combinada, chamada então de

síndrome Kwashiorkor-marasmática. O Marasmus é caracterizado por perda de

massa corpórea, particularmente muscular e gordura subcutânea e é usualmente

resultado de severa restrição de ingestão calórica. O Kwashiorkor é caracterizado por

edema, sendo resultado da deficiência prolongada da ingestão proteica e afeta

principalmente crianças (DE ANGELIS, 1986; WHO, 2011).

A DP pode afetar todos os sistemas e órgãos (TROWELL, 19541 apud

(MONTE, 2000), visto que as proteínas constituem o principal componente estrutural

celular (IMNA, 2005). Entretanto, os tecidos que apresentam alta taxa de renovação

e proliferação celular, e que, portanto, requerem um maior aporte de nutrientes, são

primeiramente afetados, como o tecido hematopoético (BORELLI et al., 2004; XAVIER

et al., 2007; BORELLI et al., 2009).

1.2 HEMATOPOESE E DESNUTRIÇÃO PROTEICA

A hematopoese é um processo hierárquico, dinâmico e finamente controlado

em que as células tronco hematopoéticas (CTH) pluripotentes se autorrenovam ou

proliferam e se diferenciam em células progenitoras para originar os diferentes tipos

celulares que compõem o sistema sanguíneo (WEISSMAN, 2000; LARSSON et al.,

2005; BRYDER et al., 2006; WEISSMAN e SHIZURU, 2008; SEITA e WEISSMAN,

1 TROWELL, H. C.; DAVIES J. N. P.; DEAN, R. F. A. Kwashiorkor. London: Edward Arnold, 1954.

15

2010). A metodologia mais utilizada para identificar e isolar as CTH e as diferentes

populações de progenitores hematopoéticos é a partir do seu imunofenótipo. Em

camundongos, essas células são comumente caracterizadas por não expressarem

marcadores de células diferenciadas (ou marcadores de linhagem, Lin).

Em camundongos, as CTH (Lin−Flk2−Thy1.1lowSca-1+c-Kit+), de origem

mesodérmica, originam uma população heterogênea de progenitores hematopoéticos

multipotentes (MPP, Lin−Flk2−Thy1.1lowSca-1+c-Kit+), com pequena ou sem

capacidade de autorrenovação. Os MPP, por sua vez, podem se diferenciar para a

linhagem linfoide, originando o progenitor linfoide comum (CLP, Lin−Il7rlowc-Kit+Sca-

1+), do qual resultam linfócitos T e B e células natural killers, e progenitor mieloide

comum (CMP, Lin−Il7r−c-Kit+Sca-1−CD34+CD16/32low), que origina os progenitores

grânulo-monocíticos (GMP, Lin−Il7r−c-Kit+Sca-1−CD34+CD16/32high) e

megacariocítico-eritroides (MEP, Lin−Il7r−c-Kit+Sca-1−CD34−CD16/32low) (Figura 1)

(KONDO et al., 1997; AKASHI et al., 1999; WEISSMAN, 2000; BRYDER et al., 2006;

WEISSMAN e SHIZURU, 2008). Estudos indicam que o comprometimento das células

tronco pluripotentes para determinada linhagem hematopoética ocorra nos estágios

iniciais da divisão celular e de forma simétrica ou assimétrica, ou seja, podendo

resultar em duas filhas com graus de comprometimento diferentes (SUDA et al., 1984;

QUESENBERRY et al., 2005).

Está bem estabelecido na literatura que a DP/DPE compromete órgãos linfo-

hematopoéticos – medula óssea (MO), baço e timo - com consequente modificação

da resposta imune (KEUSCH, 2003a; FOCK, BLATT, et al., 2010; FOCK, ROGERO,

et al., 2010; SCRIMSHAW, 2010; FOCK et al., 2012). Isto é evidenciado pela redução

da migração celular, fagocitose, atividade bactericida e fungicida e alteração na

produção de espécies reativas de oxigênio (CHANDRA, 1991; KEUSCH, 1994;

BORELLI et al., 1995; VITURI et al., 2000; NARDINELLI e BORELLI, 2001), nitrogênio

e diminuição na síntese de fator de necrose tissular (TNF) -a e interleucinas (IL) -1a,

-1β e -6 (FOCK et al., 2007).

Adicionalmente, a DP causa alterações hematológicas quantitativas na série

eritrocitária, provocando anemia não ferropriva, com redução de reticulócitos e baixa

responsividade à eritropoietina (DE ANGELIS, 1986; ROBBINS et al., 2003; BORELLI

et al., 2007; BORELLI et al., 2009). Este quadro anêmico provém da redução da

produção de células e precursores eritroides, decorrente de alterações qualitativas e

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quantitativas das células tronco e progenitoras hematopoéticas, originadas por

alterações no ciclo celular (BORELLI et al., 2007; BORELLI et al., 2009). Estas

alterações incluem aumento de células tronco e progenitoras hematopoéticas nas

fases G0/G1 do ciclo celular, bem como aumento de proteínas inibitórias da progressão

do ciclo (p21 e p27), além de redução de proteínas indutoras, como Cdk2, Cdk4,

PCNA e ciclinas D1 e E (BORELLI et al., 2009; NAKAJIMA et al., 2014).

Figura 1. Modelo hierárquico da hematopoese e caracterização fenotípica das CTH e

progenitores hematopoéticos. Os marcadores de superfície utilizados para isolamento estão

indicados à esquerda para humanos (superior) e camundongos (inferior) para cada célula

tronco ou progenitora (WEISSMAN e SHIZURU, 2008).

A DP também provoca alterações histológicas na matriz extracelular (MEC)

medular, com atrofia dos compartimentos eritroide e grânulo-monocítico (XAVIER et

<< Prev Figure 1 Next >>PMC full text: Blood. 2008 Nov 1; 112(9): 3543–3553.doi: 10.1182/blood-2008-08-078220Copyright/License ▼ Request permission to reuse

Copyright © 2008 by The American Society of Hematology

Figure 1

Schematic of hematopoietic development indicating intermediates in the hierarchy of hematopoietic differentiation. Surface markersused for isolation are indicated at left for human (top) and mouse (bottom) for each stem and progenitor cell. HSC indicates long-term

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al., 2007) e aumento do depósito de proteínas, principalmente fibronectina,

trombospondina e laminina (VITURI et al., 2000). A MEC fornece o suporte físico para

as células hematopoéticas, mas também desempenha um papel essencial na

modulação da resposta a fatores de crescimento, citocinas, hormônios e vitaminas e

desta forma, pode modular funções biológicas, como a adesão, proliferação,

diferenciação e migração das células hematopoéticas (LYRA et al., 1993; KLEIN,

1995; VITURI et al., 2000). Desta forma, alterações na MEC podem ser significantes

para a fisiologia do tecido medular, comprometendo a homeostasia do microambiente

medular (KLEIN, 1995; XAVIER et al., 2007; BORELLI et al., 2009).

1.3 MICROAMBIENTE MEDULAR E NICHO HEMATOPOÉTICO PERIVASCULAR

Em 1978, Schofield demonstrou que o microambiente medular atua como o

principal regulador da CTH, modulando seus processos de diferenciação, proliferação

e autorrenovação (SCHOFIELD, 1978).

O microambiente medular é altamente organizado e constituído basicamente

de células hematopoéticas nos mais variados estágios de maturação e pelo estroma

medular (DEANS e MOSELEY, 2000). O estroma apresenta-se como uma estrutura

compartimentalizada e dinâmica que, além de fornecer o parênquima de sustentação

celular, permite um "ambiente bioquímico" fundamental para a proliferação,

diferenciação e maturação das células hematopoéticas (MAYANI et al., 1992).

O estroma é composto pela MEC, diversas substâncias solúveis e pelo

sistema celular estromal, formado por células mesenquimais, fibroblastos, células

endoteliais, células reticulares e adipócitos, além de outros tipos celulares. O estroma

é altamente adaptativo, visto que apresenta a habilidade de alterar sua composição e

função em resposta a estímulos externos e, desta forma, modular a sobrevivência,

proliferação e o desenvolvimento das células hematopoéticas em todos os seus níveis

de diferenciação (BORDIGNON et al., 1999; ZHANG et al., 2003; KOLF et al., 2007;

SATO et al., 2010), através da produção local de citocinas e proteínas da MEC

(GORDON, 1988).

Dessa maneira, a existência de fatores regulatórios pode formar

microambientes indutivos (TRENTIN, 1978; TESTA e DEXTER, 1990), que controlam

a hematopoese pela produção e secreção local de citocinas pelas células do estroma,

18

co-localização de citocinas para a CTH nos locais de contato célula - célula e/ou célula

– MEC ou ainda, por estímulo direto pelo contato celular (RIOS e WILLIAMS, 1990;

METCALF, 1994).

O termo nicho hematopoético é utilizado para designar a localização anatômica

do microambiente em que as CTH residem (SCHOFIELD, 1978). Embora as CTH

estejam bem caracterizadas, seu nicho ainda é pouco compreendido.

A interface entre a MO e os ossos trabeculares é chamada endósteo, no qual

estão presentes numerosos osteoblastos. A primeira evidência que mostrou que os

osteoblastos poderiam modular a CTH foi um estudo de Taichman & Emerson em

1994, no qual foi demonstrado que osteoblastos diferenciados in vitro a partir da CTM

produzem fator estimulador de colônias grânulocíticas (G-CSF) (TAICHMAN e

EMERSON, 1994). Posteriormente, foi demonstrado que um grande número de CTH

situa-se próximo ao endósteo e surgiu o conceito do nicho endosteal como principal

nicho hematopoético (ZHANG et al., 2003; LI e XIE, 2005). Diversos estudos in vitro e

in vivo relatam que os osteoblastos parecem ser necessários para a manutenção da

hematopoese (KIEL e MORRISON, 2008; MENDEZ-FERRER et al., 2010;

RENSTROM et al., 2010), de forma que estas células podem regular o número e a

função das CTH através, por exemplo, da secreção de osteopontina (STIER et al.,

2005) e da expressão de angiopoietina (Ang) -1 (ARAI et al., 2004), que mantem a

CTH em estado de quiescência.

Entretanto, estudos mais recentes mostram que a modulação das CTH pelos

osteoblastos é prioritariamente indireta. Isto foi evidenciado por estudos de imagens

in vivo que demonstraram que apenas um pequeno número de CTH está em contato

com os osteoblastos (KIEL et al., 2005; KIEL et al., 2009) Além disso, em estudos

baseados na depleção ou aumento do número de osteoblastos, não foram observadas

alterações importantes nas CTH (KIEL et al., 2007; LYMPERI et al., 2008).

Duas observações importantes questionaram a existência de outros nichos

hematopoéticos: (a) durante o desenvolvimento fetal, ou seja, antes da formação das

cavidades medulares, a CTH tem capacidade de diferenciação e auto-renovação

(TAVIAN e PEAULT, 2005) e (b) a CTH pode residir próxima aos sinusóides

medulares (KIEL et al., 2005; KIEL et al., 2007; NOMBELA-ARRIETA et al., 2013).

Em 1997 foi demonstrado por Ohneda & Bautch que células endoteliais (CE)

19

sinusoidais podem modular e sustentar a CTH in vitro (OHNEDA e BAUTCH, 1997),

mas apenas em 2005, no estudo realizado por Kiel e colaboradores, foi apontada a

existência de um nicho perivascular (KIEL et al., 2005). O nicho perivascular se

localiza na região anatômica próxima ao endotélio vascular dos sinos medulares e

parece ser a localização in vivo de dois terços das CTH (MITSIADIS et al., 2007;

CARRION et al., 2010), de maneira que as CTH se mantêm próximas a CE e células

perivasculares, como a célula tronco mesenquimal (CTM) (KIEL e MORRISON, 2008;

MENDEZ-FERRER et al., 2010).

As CE sinusoidais medulares podem modular as CTH, através da expressão

de proteínas reguladoras, como proteína celular de adesão vascular 1 (VCAM-1),

CXCL-12, Ang-1 (LEVESQUE e WINKLER, 2011) e receptor de fator de crescimento

endotélio-vascular (VEGFR) - 2 (HOOPER et al., 2009). Além disso, promovem a

expansão das CTH in vitro (BUTLER et al., 2010), sugerindo que as CE podem ser

essenciais para a proliferação das CTH in vivo (WINKLER et al., 2010).

Um dos mecanismos propostos para tal é a capacidade de síntese in vitro de

stem cell factor (SCF) e fator estimulador de colônias de granulócitos e macrófagos

(GM-CSF) pelas CE, que promovem a proliferação de progenitores hematopoéticos

(WADHWA e THORPE, 2008). Entretanto outros componentes do nicho perivascular

produzem estes mediadores, como as CTM. Dessa forma, não é claro se a modulação

in vivo da CE sobre a CTH é direta ou indireta.

Na MO, a célula tronco mesenquimal (CTM) é um importante componente do

nicho hematopoético. As CTM, são um grupo de células clonogênicas presentes ao

longo da MO que originam o estroma de suporte para a hematopoese (KASSEM e

ABDALLAH, 2008; HOCKING e GIBRAN, 2010).

A primeira evidência concreta de que a MO contém células precursoras não-

hematopoéticas provém de estudos realizados por Friedenstein na década de 70

(FRIEDENSTEIN et al., 1976). Posteriormente, diversos estudos estabeleceram que

essas células exibiam capacidade de se diferenciar em tipos celulares mesodérmicos,

como osteoblastos, condrócitos e adipócitos (BARRY e MURPHY, 2004; JUNG et al.,

2009; WATABE e MIYAZONO, 2009; KURODA et al., 2010). As CTM representam

uma pequena fração (0,001-0,01%) do total de células nucleadas na MO, porém

podem ser isoladas e expandidas com alta eficiência (BARRY e MURPHY, 2004), pois

apresentam aderência seletiva a superfícies plásticas quando comparadas com as

20

células hematopoéticas. Apresentam características fusiformes e tipo fibroblastóides

e, no início do crescimento in vitro, formam colônias análogas às unidades formadoras

de colônias de fibroblastos.

As CTM são capazes de influenciar a função de outras células através de

interação direta célula-célula e através da liberação de um amplo espectro de fatores

bioativos, como citocinas e fatores de crescimento (OLIVEIRA, 2010). Como exemplo,

as CTM produzem CXCL-12 e Ang-1, que contribuem para manter a CTH quiescente,

impedindo sua proliferação e apoptose (ARAI et al., 2004). Outro regulador negativo

da hematopoese sintetizado pelas CTM é o TGF-β, que age de forma direta, inibindo

o ciclo celular de progenitores hematopoéticos, e indireta, promovendo maior

diferenciação osteoblástica da CTM (RUSCETTI et al., 2005). Em contrapartida, o

TGF-β estimula a expressão de IL-11 pelas células do estroma, que estimula a

proliferação de progenitores hematopoéticos (PAUL et al., 1990). Dados prévios do

nosso grupo mostraram que CTM de camundongos desnutridos sintetizam menor

quantidade de CXCL-12 e maior quantidade de SCF que camundongos bem nutridos

(HASTREITER, 2014).

Além disso, alguns subtipos de CTM exercem atividade essencial no nicho

perivascular. Foi demonstrado que CTM que expressam nestina (Nes+), receptor de

leptina (LepR+) e “CAR cells” – células reticulares que sintetizam grandes quantidades

de CXCL-12 - se localizam adjacentes às CE e às CTH, sendo que estas células

expressam fatores de crescimento e citocinas que favorecem a manutenção e

proliferação das CTH, como SCF e CXCL-12 (SUGIYAMA et al., 2006; MENDEZ-

FERRER et al., 2010; EHNINGER e TRUMPP, 2011; DING et al., 2012). Entretanto,

a distribuição destas células não é uniforme. Kunisaki e colaboradores apontaram que

CTM Nes+/NG2-/LepR+ estão localizadas no nicho perivascular sinusoidal e CTM

Nes+/NG2+/LepR- se encontram preferencialmente no nicho perivascular arteriolar,

promovendo a manutenção da quiescência das CTH (KUNISAKI et al., 2013) (Figura 2).

21

Figura 2. Ilustração dos nichos hematopoéticos perivasculares (BOULAIS e FRENETTE,

2015).

Além de exercerem um papel regulador sobre as CTH, as CTM modulam

células da linhagem endotelial, de maneira que é cogitado que a função primária das

CTM é fornecer suporte para a hematopoese e estabilizar os vasos sanguíneos

medulares (BIANCO et al., 2010). A habilidade de diferenciação das CTM para células

endoteliais tem sido amplamente utilizada na literatura, entretanto os mecanismos e

as moléculas envolvidas nestes processos não estão totalmente esclarecidos

(OSWALD et al., 2004; LIU et al., 2007; LOZITO, KUO, et al., 2009; LOZITO, TABOAS,

et al., 2009; MOSNA et al., 2010). Em trabalho anterior do nosso grupo, mostramos

que as CTM podem se diferenciar em CE, tanto em camundongos nutridos quanto em

desnutridos, através da cultura com meio de crescimento endotelial, visto que

adquirem características de células endoteliais (HASTREITER, 2014). Entretanto,

como o fenótipo dessa célula diferenciada não é idêntico ao das células endoteliais,

alguns autores as intitulam como célula endotelial “like” ou endotélio ”like” (LIU et al.,

2007).

Desde a descoberta do nicho perivascular, foi proposto que as CTH com

proliferação mais ativa se situavam preferencialmente na região perivascular,

22

enquanto que as CTH quiescentes preferencialmente situavam-se no nicho endosteal,

sob influência osteoblástica (LEVESQUE e WINKLER, 2011). Atualmente, os estudos

mostram que a regulação sobre a hematopoese é predominantemente do nicho

perivascular, tanto arteriolar quanto sinusoidal. Contudo, a real existência in vivo

destes nichos e seus componentes precisa ser melhor elucidada.

23

2 HIPÓTESE E OBJETIVOS

A desnutrição proteica compromete a hematopoese, tanto quantitativamente

quanto qualitativamente. Entretanto, pouco se sabe sobre os mecanismos envolvidos

na regulação da hematopoese numa situação de desnutrição proteica.

Sabendo que o nicho perivascular é um importante regulador da hematopoese

e que a desnutrição proteica altera a função das células tronco mesenquimais e das

células endoteliais, nossa hipótese é que a desnutrição proteica tem uma ação

fisiopatológica importante, comprometendo mecanismos essenciais de controle da

hematopoese. Sendo assim, um possível comprometimento do microambiente

hematopoético pode ser um dos mecanismos que levam à hipoplasia medular

observada na desnutrição.

Portanto, esta pesquisa teve como objetivo averiguar os efeitos da

desnutrição proteica nas células tronco mesenquimais e nas células endoteliais e

verificar as consequências destas alterações sobre a proliferação e a diferenciação

das células tronco e progenitoras hematopoéticas in vitro. Além disso, objetivamos

avaliar o efeito da desnutrição proteica sobre alguns aspectos intrínsecos das células

tronco e progenitoras hematopoéticas relacionados à granulopoese.

24

3 CAPÍTULO I

A DESNUTRIÇÃO PROTEICA SUPRIME A HEMATOPOESE ATRAVÉS DO

COMPROMETIMENTO DAS CÉLULAS ENDOTELIAIS MEDULARES

25

Title: Protein malnutrition halts hematopoiesis by bone marrow endothelial impairment

Authors: Araceli Aparecida Hastreiter1, Guilherme Galvão dos Santos1, Ed Wilson

Cavalcante Santos1, Edson Naoto Makiyama1, Primavera Borelli1, Ricardo Ambrósio

Fock1*

1 Department of Clinical and Toxicological Analysis, School of Pharmaceutical

Sciences, University of São Paulo, São Paulo, Brazil.

* To whom correspondence should be addressed. Fock, Ricardo Ambrósio. Laboratory

of Experimental Hematology, Department of Clinical and Toxicological Analysis,

School of Pharmaceutical Sciences, University of São Paulo. Avenida Lineu Prestes,

580 - Bloco 17. São Paulo, SP, Brazil. 05508-900. Phone: +551130913639. e-mail:

[email protected]

26

ABSTRACT

Protein malnutrition (PM) affects tissues with high rate of cell renewal and proliferation,

such as the hematopoietic system. PM affects hematopoiesis leading to bone marrow

(BM) hypoplasia and arrests hematopoietic stem cells (HSC) in G0/G1 cell cycle

phases, which cause anemia and leukopenia. HSC possess the ability to differentiate

into all functional blood cells as well as to self-renewal without differentiation. In this

context, hematopoiesis is mainly regulated by BM niches where endothelial cells (EC)

present a key regulatory role. In this study, we assessed the impact of PM on

hematopoietic stem and progenitor cells and the role of BM endothelial cells upon

hematopoietic impairment in a murine model. We showed that PM decreases HSC and

hematopoietic progenitor pool, in addition to the inability of the BM of malnourished

animals to sustain hematopoiesis. Furthermore, PM committed hematopoietic

regulatory characteristics from EC, resulting in the modification of both cell cycle

pattern and hematopoietic differentiation. Thus, since PM disturbs EC, it become one

of the factors responsible for the hematopoietic cell cycle arrest and impairment of HSC

differentiation.

Key-words: Protein malnutrition; Hematopoietic stem cell; Endothelial cell;

Hematopoiesis regulation.

27

Abbreviations ANG Angiopoietin BM Bone marrow CAR CXCL-12 abundant reticular cells CLP Common lymphoid progenitor CXCL-12 C-X-C motif chemokine 12 DMEM Dulbecco’s Eagle modified medium EC Endothelial cell EDTA Ethylenediamine tetraacetic acid EGM Endothelial cells growth medium ELISA Enzyme-linked immunosorbent assay G-CSF Granulocyte colony stimulating factor GM-CSF Granulocyte and macrophage colony stimulating factor GMP Granule-monocytic progenitor HSC Hematopoietic stem cell IL Interleukin KO Knock out LEPR Leptin receptor LSK Lineage, Sca-1 and c-Kit negative cell MEP Megakaryocytic-erythroid progenitor MNC Mononuclear cell MPP Multipotent progenitor MSC Mesenchymal stem cell PBS Phosphate-buffered saline PM Protein malnutrition qPCR Quantitative polymerase chain reaction SCF Stem cell factor VCAM Vascular cell adhesion protein

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INTRODUCTION

Malnutrition has been put aside for many decades but rises again due to an

increase in elderly and chronic disease patients, as well as hunger driven by conflict

and climate change (FAO, 2018). Protein malnutrition (PM) is the most common type

of malnutrition and leads to several physiological consequences, depending on its

duration and intensity (FAO, 2012; FAO, 2018).

Hematopoietic tissues are the ones to be firstly affected by PM due to their

continuous turnover (BORELLI et al., 2004; XAVIER et al., 2007; BORELLI et al., 2009;

SANTOS et al., 2017). PM yields alterations in lymphohematopoietic organs (bone

marrow (BM), spleen, and thymus), resulting in anemia, leukopenia, and alterations in

the immune response, increasing susceptibility to infections (KEUSCH e FARTHING,

1986; BORELLI et al., 1995; VITURI et al., 2000; KEUSCH, 2003b; FOCK et al., 2007).

Hematopoiesis is a dynamic process in which all mature blood cells are formed

from a pluripotent hematopoietic stem cells (HSC). HSC have the ability to self-renew

and differentiate into hematopoietic multipotent progenitors (MPP), hierarchically

giving rise to lineage-specific progenitors – common lymphoid (CLP) and common

myeloid progenitors (CMP). CMP can differentiate into granule-monocytic (GMP) or

megakaryocytic-erythroid (MEP) progenitors, which father all myeloid mature cells

(WEISSMAN e SHIZURU, 2008). This whole process is meticulously controlled by

hematopoietic transcription factors that are activated by various cytokines and growth

factors such as SCF, IL-11, IL-3, G-CSF, GM-CSF, and others, to maintain an

adequate balance between self-renewal and differentiation to preserve HSC pool. The

source of the vast majority of signals modulating hematopoiesis and the location where

HSC reside in the bone marrow (BM) is named hematopoietic niche (SCHOFIELD,

1978). The hematopoietic niche supports all stages of hematopoiesis, from the self-

renewal of HSC to the release of mature cells into the peripheral blood (MAYANI et al.,

1992).

There is poor evidence regarding the effects of PM on hematopoietic niches.

For a long time, it was believed that the main modulator of hematopoiesis was the

endosteal niche. Several studies have shown how osteoblasts regulate the HSC pool

in vivo (ARAI et al., 2004; ARAI e SUDA, 2007; EMA e SUDA, 2012), but with the

improvement of technical analysis and supplies and the use of KO mice, few effects

on HSC were actually attributed to endosteal niche (MENDEZ-FERRER et al., 2010;

29

KUNISAKI et al., 2013).

On the other hand, recent studies report that the perivascular niche is the real

modulator of hematopoiesis (MENDEZ-FERRER et al., 2010; KUNISAKI et al., 2013;

BOULAIS e FRENETTE, 2015; XU et al., 2018), which was first described in 2005 and

is located near the vascular endothelium of the bone marrow sinusoids and arterioles

(KIEL et al., 2005; MITSIADIS et al., 2007; CARRION et al., 2010). In addition to

mesenchymal stem cells (MSC), endothelial cells (EC) are a critical component of the

normal hematopoietic niche and display an HSC-supportive activity (LI et al., 2010).

Bone marrow EC promote HSC maintenance by SCF production (DING et al., 2012;

XU et al., 2018), regulate the mobilization of hematopoietic cells by CXCL-12,

angiopoietin and VCAM-1 production (LEVESQUE e WINKLER, 2011) and are the

mainly endogenous producer of G-CSF, which engage granulopoiesis.

Although hematopoietic changes caused by PM have been described for a long

time, little information is known concerning the alterations in HSC, primarily in cell

differentiation, as only a few mechanisms are described so far. Likewise, there is no

literature available covering the effects of PM on perivascular niche. Thus, this study

aims to provide novel information concerning the alterations in bone marrow EC and

its regulatory function caused by PM.

RESULTS Protein malnutrition decreases hematopoietic stem and progenitor cells in bone

marrow and leads to anemia and leukopenia

In the present study, we used a low-protein diet to induce protein malnutrition

and evaluated the hematologic consequences in murine. Mice from malnourished and

control groups exhibited similar food intake during the period of malnutrition induction,

however the malnourished group (PM group) had lower protein intake due to

hypoproteic diet. As consequence, malnourished mice presented body weight loss and

decreased serum protein and albumin concentrations (Table 1).

In addition, mice that received hypoproteic diet showed quantitative alterations

in the erythroid parameters of peripheral blood, with reduction in erythrocyte count,

hemoglobin concentration and hematocrit values. The PM group presented expressive

leukopenia with decreased number of neutrophils, lymphocytes, and monocytes

30

(Table 1). However, no significant morphological differences in cells between groups

were found. The PM group also showed a hypoplastic bone marrow and a significant

reduction in the total nucleated cell count and in the absolute value of all lineages

(Table 1).

Since protein malnutrition caused medullary hypoplasia and decreased number

of blasts, the different populations of hematopoietic progenitors were quantified by flow

cytometry to investigate whether the changes were progenitor-specific. PM group

showed a decrease in HSC (Lin−Flk2−Thy1.1lowSca-1+c-Kit+) population (Fig. 1a–c) as

well as in MPP (Lin−Flk2−Thy1.1lowSca-1−c-Kit+), CLP (Lin−Thy1.1−Il7r+c-Kit+Sca-1+),

CMP (Lin−Il7r−c-Kit+Sca-1−CD34+CD16/32low), GMP (Lin−Il7r−c-Kit+Sca-

1−CD34+CD16/32high), and MEP (Lin−Il7r−c-Kit+Sca-1−CD34−CD16/32low) populations

(Fig. 1a–c), indicating that PM affects the number of all myeloid and lymphoid

progenitors.

To understand why there is reduction of both HSC and hematopoietic progenitor

cells in the PM group, the expression of transcription factors that control pluripotency

and guide cell differentiation was quantified in bone marrow c-Kit+ population isolated

from BM. The expression of the pluripotency transcription factors Sox-2, Nanog, and

Pou5f1 (Oct-4) was impaired in malnourished mice (Fig. 1d), which indicates that PM

can imbalance HSC self-renewal and differentiation. Additionally, the PM group

presented a higher percentage of MNC in the G0/G1 phases (Fig. 1e–h), whereas no

differences in viable and apoptotic cells were observed (Fig. 1i and 1j). As the activity of the differentiation transcription factors often overlaps and

fluctuates in the various types of hematopoietic progenitors (ZHU e EMERSON, 2002;

MONTICELLI e NATOLI, 2017), here they were classified in promoters of myeloid or

lymphoid differentiation, according to their most prominent activity. The PM group

exhibited a decreased expression in c-Kit+ cells of myeloid factors Gata1, Gata2, Nfe2,

Spi1, and Cebpa, as well as lymphoid factors Gata3 and Ikzf3 (Fig. 1d), denoting that

PM prevents cellular differentiation, likewise self-renewal.

Protein malnutrition impairs proliferation and delay cell cycle of leukemic cells

To elucidate if the alterations observed in PM are intrinsic to HSC and

hematopoietic progenitors or if they are due to alterations in the hematopoietic niche,

we investigated whether the BM of PM group supports hematopoiesis through cell

31

cycle evaluation of highly proliferative transplanted hematopoietic cells. Syngeneic

transplants with labeled C1498 leukemic cell lineage were performed (Fig. 2a). After 4

days of transplantation, no C1498 cells were found in the peripheral blood or spleen of

both groups, but more were found in the BM of malnourished mice in comparison to

control mice (Fig. 2b-d). However, in the PM group, transplanted C1498 cells showed

increased numbers of cells in G0/G1 cell cycle phases (Fig. 2e–g), as well as reduced

percentages in the S phase and lower proliferation index (Fig. 2h and 2i). The cell

cycle arrest observed in the transplanted C1498 cells is similar to that observed in the

ex vivo hematopoietic stem and progenitor cells in the PM group, demonstrating that

PM implies hematopoietic extrinsic changes that can lead to bone marrow hypoplasia.

Protein malnutrition does not impair mesenchymal-to-endothelial transdifferentiation

As the vascular niche is an important regulator of HSC in vivo (MENDEZ-

FERRER et al., 2010; KUNISAKI et al., 2013; BOULAIS e FRENETTE, 2015), we

investigated whether alterations on BM-EC could be one of the mechanisms that leads

to the hypoplasia observed in the PM group. Therefore, BM-MSC were collected from

control and PM groups, expanded to passage 2-3, and transdifferentiated to EC. BM-

MSC from both groups were able to differentiate into adipocytes and osteocytes after

cultivation in respective media (Fig. 3a–e), and showed typical immunophenotypic

labeling (CD90.1+CD49e+CD44+CD34−CD45−CD11b−) with low positivity for cells

marked with Sca-1 and CD105 (Fig. 3f). No differences were observed in MSC

phenotype between control and malnourished groups.

First, we investigated whether PM impairs mesenchymal-to-endothelial

transdifferentiation by morphological analysis, flow cytometry, and qPCR techniques.

Before the endothelial differentiation, cells presented polygonal and spindle shapes,

which are characteristics of MSC (Fig. 3g and 3h). After the differentiation, cells

acquired a sharp morphology (Fig. 3i) and formed structures similar to microtubules in

Matrigel® (Fig. 3j). No quantitative or morphological differences were observed in the

differentiation pattern between control and PM groups.

After the endothelial differentiation, CD31, CD144, and Sca-1 became positive

(Fig. 3f) and were similar in both groups. Gene expression screening on endothelial

cells reinforced these results, through an increase of 2-8-fold-change on the

endothelial characteristic genes Flt1 (VEGFR1), Kdr (VEGFR2), Vcam1, and Nt5e

32

(CD73) (Fig. 3k-n) with no difference between control and PM groups in these

parameters. Additionally, the gene expression of Mcam (CD146), Pdgfb1, Nes (nestin),

Cspg4 (NG2), and Lepr (leptin receptor) was evaluated, which are important

perivascular mesenchymal markers in the BM niche. EC lacked Mcam and Pdgfb1

(Fig. 3o and p) and enhanced Lepr (Fig. 3q) expressions in both groups, but no

difference in Nes and Cspg4 expressions were detected (Fig. 3r and s) between

groups.

Protein malnutrition affects the function of endothelial cells

To evaluate the hematopoietic modulatory properties of EC, the production of

hematopoietic regulatory cytokines Ang-1, SCF, CXCL-12, IL-11, TGF-b, GM-CSF,

and G-CSF was quantified on the supernatant of EC cultures from both groups.

Additionally, the expression of genes related to HSC maintenance and hematopoietic

progenitor/precursor differentiation was also evaluated by qPCR (Fig. 4a). EC from

both groups produced large amounts of Ang-1, however, cells from the PM group

presented a significantly lower amount of Ang-1 (Fig. 4b) in comparison to cells from

the control group. SCF and CXCL-12 production were also decreased in cells from the

PM group (Fig. 4c and 4d), and their respective gene expression was downregulated

(Kitl and Cxcl12) (Fig. 4a). Concerning the regulation on the capacity of differentiation,

EC from PM mice presented an increase in gene expression and production of IL-11

(Fig. 4a and 4e), but none difference in production of G-CSF (Fig. 4h) despite the Csf3

upregulation observed in cells obtained from the PM group (Fig. 4a). Alterations on

TGF-b levels was not observed between groups (Fig. 4f and 4g), and GM-CSF was

not detected in both control and PM groups. Additionally, IL-3 was also not detected,

neither by ELISA nor Il3 gene expression quantification by qPCR.

PM shifts hematopoietic differentiation via EC

Since PM causes anemia and leucopenia associated with BM hypoplasia and

changes the synthesis of hematopoietic mediators of differentiation by EC, such as IL-

11 and G-CSF, the ability of EC to induce hematopoietic differentiation was analyzed

in a culture system with and without cell contact. First, EC from both groups were cross-

cultured with c-Kit+ cells, and the differentiation line from CMP to granulocytic cells was

33

evaluated. After 3 days of co-culture, the number of c-Kit+ cells from the PM group was

higher when cultured with control EC compared to c-Kit+ cells from the control group

cultured with control EC. c-Kit+ cells from the PM group cultured with malnourished EC

did not show differences among groups (Fig. 5a). Interestingly, the PM group

presented more CMP than observed in the control group even though EC did not affect

the percentage of CMP (Fig. 5b). Additionally, EC affects hematopoietic differentiation

(Fig. 5c-e). Malnourished EC promoted increased differentiation to MEP (Fig. 5d) but

decreased differentiation to GMP (Fig. 5c) and granulocytes (Fig. 5e). About the

macrophage quantification, no differences were observed among groups (Fig. 5f). For a better understanding of the paracrine effects of EC, cultures conditioned

with EC supernatant of both groups were performed with BM-MNC for cellular

population evaluation (Fig 5g-l), as well as with c-Kit+ cells for gene expression

analysis (Fig. 5m-o). HSC quantification was reduced in cultures performed with

malnourished EC supernatant compared to control EC supernatant (Fig. 5g), in

agreement with decreased SCF production and low Gata2 expression (Fig. 5n), a

hematopoietic self-renewal transcription factor. The expression of the pluripotency

genes Sox2 and Nanog were also downregulated in cells from both groups cultured

with malnourished EC supernatant compared to cells cultured with control EC

supernatant (Fig. 5m). These results may justify a repercussion of the decreased

number of HSC observed in the culture system of cells with malnourished EC

supernatant.

Additionally, although neither malnourished EC nor control EC supernatants

altered the MPP quantification, the PM group showed higher percentage of HSC in

comparison to the control group (Fig. 5h). However, cells from the PM group cultured

with control EC supernatant showed higher values for CMP and GMP (Fig. 5j and 5k)

in comparison to malnourished cells cultured with malnourished EC supernatant. About

the MEP quantification, the PM group showed higher percentage of MEP in

comparison to the control group, but the EC supernatant from both groups did not affect

these results (Fig. 5l). Regarding the lymphoid differentiation, EC seemed to have no influence in

malnourished cells (Fig. 5i). We observed a very reduced number of CLP after the

conditioned cultures with malnourished EC supernatants (Fig. 5i), and c-Kit+ cells did

not express the transcription factors that regulate the lymphoid differentiation (Gata3

and Ikzf3). The quantification of CMP, GMP, and MEP in co-cultures and conditioned

34

cultures agreed among themselves, but GMP values in co-culture were up to 3 times

higher than in the conditioned culture (Fig. 5j–l). Accordingly, PM modulated Sfpi1 and Cebpa expressions, which are

transcription factors of the beginning and end of granulocyte differentiation,

respectively (Fig. 5n). Gata1 and Nfe2, the most important transcriptions factors for

erythroid and megakaryocytic differentiation, were upregulated in malnourished

conditioned cultures, exhibiting paracrine effects of EC. However, the co-cultures had

MEP values up to 45 times higher than in the conditioned culture, indicating that cell-

EC contact is important for granulocytic differentiation and even more for

megakaryocytic and erythrocytic differentiation. Additionally, EC did not affect the gene

expression of Il3ra in c-Kit+ cells, but c-Kit+ cells from malnourished animals cultured

with malnourished EC showed increased expression of Cxcr4 (Fig. 5o).

Endothelial cells halt cell cycle in protein malnutrition

Since malnourished EC produced less amount of Ang-1, SCF, and CXCL-12

than control EC (Fig. 4b–d), the effect of EC supernatants on the viability and cell cycle

of hematopoietic cells was investigated. BM-MNC from both groups were cultured with

EC supernatant from the control or malnourished group, and the viability, apoptosis

status, and cell cycle were evaluated by flow cytometry. The first important point is that

the cultures performed with conditioned media were able to maintain cell viability as

well as avoid apoptosis than the cultures performed with culture medium alone (Fig. 6a and 6b). Moreover, the conditioned cultures with EC supernatant induced

quantitative alterations in cell cycle phases. Cells from both groups were more frequent

in G0/G1 cell cycle phases when cultured with malnourished EC conditioned media

and, consequently, less frequent in S/G2/M cell cycle phases (Fig. 6c–h).

The effects on cell cycle entailed by malnourished EC supernatant were similar

to the changes caused by PM observed ex vivo, which strongly indicate that the

mechanism for cell cycle arrest is intrinsically related to EC. To confirm this evidence,

we performed EC conditioned cultures with a highly proliferative cell line. Leukemic

C1498 cells were cultured with EC supernatant for evaluation of cell cycle regulatory

genes by qPCR, and the results showed that malnourished EC supernatant was able

to downregulate the gene expression of the cell cycle promoters Ccnd1 (cyclin D1) and

35

Ccne1 (cyclin E1) and upregulate their inhibitors Cdkn1a (p21) and Cdkn1b (p27) (Fig. 6i–l).

DISCUSSION

The entire hematological consequences of PM remain unknown. Many studies

describe alterations in the peripheral blood, but the mechanism of how these occur and

the alterations in the BM are scarce. In this study, mice fed with hypoproteic diet (2%

protein) presented quantitative reduction of hematopoietic cells in both peripheral and

central compartments. Malnourished mice were leukopenic, which was reflected by a

lower absolute value of circulating lymphocytes, neutrophils, and monocytes in the

peripheral blood. This leukopenia can compromise both the innate and acquired

immunity, as described in previous studies (FOCK et al., 2007; FOCK, BLATT, et al.,

2010; FOCK, ROGERO, et al., 2010).

Previous report showed a decreased Lin-Sca-1+c-Kit+ (LSK) and CD45+CD34+

populations caused by PM (BORELLI et al., 2009; NAKAJIMA et al., 2014) but both

LSK and CD45+CD34+ represent heterogeneous cellular populations that include HSC

and some hematopoietic progenitors. This is the first report that shows in more detail

that all hematopoietic progenitors (MPP, CLP, CMP, GMP, and MEP) and HSC are

reduced in PM. Thus, PM does not affect any specific progenitor but all young cells in

BM. We observed that the decrease in HSC and progenitor quantification ex vivo is not

due to apoptotic process increments but is caused by cell cycle arrest, in agreement

with a previous study that reported a higher percentage of LSK cells in the G0/G1

phases in PM (BORELLI et al., 2007). PM suppressed the expression of the

pluripotency genes Sox2, Pouf51, and Nanog, thus compromising HSC self-renewal

and ability to recover the hematopoietic tissue. Also, this lower expression may be due

to the lower frequency of HSC in the malnourished group. This is a limiting factor of

the present study, however, working with a pure HSC population presented technical

difficulties due to the low frequency of this cell in BM, especially in the malnourished

group.

We demonstrated that PM decreases similarly CLP, CMP, MEP, and GMP,

corroborating hemogram and myelogram results. This lack is a consequence of the

downregulation on the expression of transcription factor genes that drive HSC and

36

progenitor differentiations in a lineage-specific manner. The main transcription factors

related to lymphoid differentiation are Gata3 and Ikzf3 and to myeloid differentiation,

they are Gata1, Gata2, Nfe2, Spi1, and Cebpa. These transcription factors do not only

act on a specific type of hematopoietic cell with their concentrations fluctuating during

hematopoiesis, but, in general, we can infer that Gata3 induces T lymphocyte

differentiation while Ikzf3 directs B lymphocyte differentiation (NAKAJIMA, 2011). On

the other hand, the myeloid transcriptions factors Gata1 and Nfe2 induce

megakaryocytic and erythroid differentiation, whereas Spi1 and Cebpa control different

stages of granulocytic differentiation (IWASAKI et al., 2006; MONTICELLI e NATOLI,

2017). Gata2 is also involved in erythroid differentiation, but more importantly, it is

related to the self-renewal capacity of HSC and MPP (IWASAKI et al., 2006). Here we

showed that PM yielded lower gene expression levels, explaining, in part, leukopenia

and anemia found in malnourished mice.

As BM niches modulate the entire hematopoietic process, we performed

syngeneic transplantation with leukemic myelo-monoblasts cell line to evaluate if

hematopoietic niches are impaired in PM. We observed a higher number of leukemic

cells in the BM of malnourished mice, but it is still necessary to investigate this major

tropism for BM, especially in BM transplantations in malnourished patients, wherein

the niche shows a key role in engraftment and chemotherapy response. Possibly, the

increased deposits of endosteal and paratrabecular fibronectin, as well as perivascular

laminin, in BM observed in PM mediate the largest number of transplanted leukemic

cells found in the BM of malnourished mice, since these proteins are important

extracellular matrix adhesion molecules for HSC and hematopoietic progenitor

retention (VITURI et al., 2000; XAVIER et al., 2007).

Previous reports showed that syngeneic transplantation distributes C1498 cells

between several tissues, such as BM, lungs, liver, spleen, and lymph nodes (MOPIN

et al., 2016). Since we did not find C1498 cells in the spleen of the animals in both the

control and malnourished groups, the time of evaluation of the mice after the transplant

is perhaps a differential factor in the cellular distribution. However, although more

leukemic cells were found in malnourished BM, these cells showed a cell cycle arrest

similar to that observed with hematopoietic cells, confirming our thoughts that

malnourished BM does not support adequately the hematopoiesis.

The perivascular niche presents a heterogeneity of cells that can modulate

hematopoiesis. The identification and role of distinct types of EC remain not completely

37

understood, but studies with different phenotypes of EC evidence their importance in

the modulation of hematopoiesis (SASINE et al., 2017; KENSWIL et al., 2018). Recent

studies suggested that arteriolar EC are the main producer of SCF and promote HSC

maintenance (XU et al., 2018), while sinusoidal EC control both hematopoietic

differentiation and the release of mature cells to peripheral blood (BOULAIS e

FRENETTE, 2015). However, the distinction between these cells is not fully

established as few in vivo reports were performed. Besides, it is still unknown whether

they act only via paracrine signs or whether cell-cell contact is indispensable.

Usually, BM-EC are defined as CD144+CD31+ cells with absence of

hematopoietic and mesenchymal markers (DING et al., 2012). In the current work, we

obtained CD144+CD31+ EC from BM-MSC transdifferentiation and disclosed that PM

did not impair this process. Although no changes in the phenotype were observed, the

function of these cells is altered by disturbance on SCF, IL-11, and G-CSF production.

The evaluated paracrine effect and contact between EC and hematopoietic cells

demonstrate that PM redirected granulocytic to megakaryocytic and erythrocytic

differentiation.

Granulopoiesis is extremely dependent on G-CSF, controlling not only

granulocytic proliferation, but also the activation and migration of mature neutrophils

by directly regulating Sfpi1 (PU.1) expression (LIESCHKE et al., 1994; LIU et al., 1996;

SEMERAD et al., 1999; KOLACZKOWSKA e KUBES, 2013). As malnourished EC had

none differences in G-CSF synthesis in vitro when compared to control EC, we infer

neutropenia observed in PM cannot be, at least in part, due to endothelial G-CSF

production. Further studies should be performed to elucidate the mechanisms by which

PM changes the function of granulocytes, whether intrinsic or caused by some other

cell in the niche.

We have shown that the participation of EC in lymphopenia observed in

malnourished mice appears to be undermost. Although CXCR-4 expression in MPP is

relevant to CLP differentiation and EC increased Cxcr4 expression in c-Kit+ cells in

vitro, rare CLP were detected in conditioned cultures, and the expression of lymphoid

transcription factors were quite downregulated. In fact, the most important promoters

of MMP to CLP differentiation are CXCL-12 abundant reticular (CAR) cells, which are

a subtype of perivascular MSC through the release of IL-7 (CORDEIRO GOMES et al.,

2016). Since hematopoiesis is a dynamic process, the fact that malnourished EC

reduced the differentiation of CLP and CMP into GMP, it itself can direct the

38

differentiation to MEP. Moreover, EC boosts IL-11 in PM, which indirectly improves

megakaryocytopoiesis and erythropoiesis, and in a lesser extent lymphopoiesis

through a synergistic effect with other cytokines and growth factors, such as IL-3, IL-

4, and SCF (WADHWA e THORPE, 2008). Even though malnourished EC conditioned

cultures improved Gata1 and Nfe2 expressions, the EC-cell contact is more important

for differentiation in MEP, but the activation pathway remains unknown.

Perhaps the most significant effect of PM in the modulation of EC over

hematopoiesis is related to the cell cycle. Malnourished EC produced less SCF than

control group, and SCF mediates HSC proliferation by direct regulation of the entry of

hematopoietic cells into the cell cycle (LENNARTSSON e RONNSTRAND, 2012).

Conditional deletion of SCF in endothelial and Lepr+ perivascular cells, but not in

osteoblasts and Nestin+ cells, leads to HSC exhaustion (DING et al., 2012).

Accordingly, a smaller amount of HSC was detected in malnourished EC conditioned

cultures. Whereas PM induced a cell cycle arrest in both hematopoietic and

transplanted C1498 cells, we evaluated the paracrine impact of EC in cell cycle

induction and inhibitory proteins. PM induces the expression of the inhibitory proteins

p21 and p27 but suppresses the induction proteins cyclin E, cyclin D1, Cdk2, Cdk4,

and Cdc25a (NAKAJIMA et al., 2014). Cyclin D1 promotes the transition from G0 to G1

cell cycle phases and is directly inhibited by p21, which keeps the cell in a quiescent

state, while cyclin E induces the progress of cell cycle from G1 to S phases. Cyclin E

is inhibited by p27, which prevents synthesis of cell mitosis (GUO et al., 2005;

PIETRAS et al., 2011). Malnourished EC downregulated cyclin E and D1 genes

(Ccne1 and Ccnd1, respectively) and upregulated p21 and p27 genes (Cdkn1a and

Cdkn1b, respectively) in leukemic cells in vitro, indicating that the quiescent induction

of HSC in PM is, at least in part, due to a cell cycle inhibitory effect of EC. In conclusion,

PM affects hematopoiesis at the hematopoietic stem and progenitor cell levels.

Furthermore, EC alterations may define hematopoietic fate under PM conditions, but

we cannot infer that the hematopoietic impairments observed are only due to EC

alterations, nor if these changes are reversible or permanent.

MATERIALS AND METHODS Mice and diets

39

All experiments were performed in accordance with the approved guidelines

by the Institutional Animal Care and this work was approved by the Animal

Experimentation Ethics Committee of the School of Pharmaceutical Sciences of the

University of São Paulo.

Male mice of the C57BL/6 inbred strains of 45- to 60-days-old were obtained

from the Production and Experimentation Laboratory of the School of Pharmaceutical

Sciences of the University of São Paulo and maintained in individual cages at 22-25°C

and relative humidity at 55 ± 10% with a regular 12-hour light/dark cycle. Mice

underwent an adaptation period (10 to 15 days) in which all animals received

normoproteic diet and water ad libitum until stabilization of body weight. After this

period, mice were divided into two groups which received either normoproteic diet

(control group) or hypoproteic diet (malnourished group).

Normoproteic and hypoproteic diets were prepared in-house. Mineral and

vitamin mixtures were prepared according to the recommendations of the American

Institute of Nutrition (AIN-93M) for adult mice (REEVES et al., 1993; REEVES, 1997).

The protein source used was casein (>85% protein) and normoproteic and hypoproteic

diets contained 12% and 2%, respectively. Both diets contained 100 g kg-1 sucrose, 80

g kg-1 soybean oil, 10 g kg-1 fiber, 2.5 g kg-1 choline bitartrate, 1.5 g kg-1 L-methionine,

40 g kg-1 of mineral mixture, and 10 g kg-1 of vitamin mixture. The control diet contained

120 g kg-1 casein and 636 g kg-1 cornstarch, while the malnourishment diet contained

20 g kg-1 casein and 736 g kg-1 cornstarch. With the exception of the protein and

cornstarch content, the two diets were identical and isocaloric, providing 1,716.3

kJ/100 g. The final protein content of both diets was confirmed by the standard micro-

Kjeldahl method.

The period for the induction of malnutrition was 35 to 40 days, and the

nutritional evaluation was performed by monitoring body weight, food consumption,

and protein intake every 48 hours during the experimental period (XAVIER et al., 2007;

DOS SANTOS et al., 2017). The variation in body weight was calculated as a relative

value between the body weight on the first day of induction to malnutrition and the last

day of this period.

Hemogram

40

Blood samples were collected with EDTA (Merck, Darmstadt, Germany) from

both control and malnourished animals. Hemograms were obtained by loading blood

samples into ABX Micros ABC Vet® equipment (Horiba ABX, Montpellier, France). The

morphological and leukocyte differential analyses were performed on blood smears

stained by May-Grünwald-Giemsa (Merck, Darmstadt, Germany) technique.

Serum protein and albumin quantification After malnourishment induction, mice were euthanized, blood samples were

collected, and the serum was separated by centrifugation (1,000 x g for 10 minutes at

4°C). The concentrations of serum proteins and albumin were determined by the use

of commercial kits (Labtest Diagnóstica SA, Lagoa Santa, Brazil) and based on

standard methods.

Myelogram

BM cells were obtained by flushing femurs with Dulbecco’s modified Eagle’s

medium (DMEM) containing low glucose (Vitrocell Embriolife, Campinas, Brazil)

supplemented with 10% fetal calf serum (Vitrocell Embriolife, Campinas, Brazil) and

0.1% penicillin and streptomycin (Sigma Aldrich, St. Louis, USA). BM cellularity was

determined by counting obtained cells using a Neubauer hemocytometer, and

myelogram was performed by morphological and differential analysis on

cytocentrifugated smears stained by May-Grünwald-Giemsa standard method.

Bone marrow mononuclear and c-Kit+ cells isolation

Total BM cells of both femurs and tibias were flushed with McCoy 5A (Sigma

Aldrich, St. Louis, USA) supplemented with 10% fetal calf serum (Vitrocell Embriolife,

Campinas, Brazil) and 0.1% penicillin and streptomycin (Sigma Aldrich, St. Louis,

USA), then Mononuclear cells (MNC) were separated by density gradient with Ficoll-

Histopaque technique (Sigma Aldrich, St. Louis, USA). After that, MNC were labeled

with anti-CD117 microbeads (Miltenyi Biotech Inc., Auburn, USA), and c-Kit+ cells were

isolated on a magnetic column following the manufacturer´s instructions.

In vivo transplantation

41

C1498 cells (TIB-49ä, ATCCâ) were labeled with PKH26 Red Fluorescent Cell

Linker Kit (Sigma Aldrich, St. Louis, USA) following the manufacturer´s instructions. 5

x 106 C1498 cells were resuspended in 150 µL of sterile and apyrogenic saline and

injected in the caudal vein of control and malnourished mice at the end of the period

of malnutrition induction. Mice were monitored every 24 hours and after 4 days, blood,

spleen, and bone marrow cells were collected for flow cytometry analysis.

Cell culture Mesenchymal stem cells isolation

MSC were obtained and characterized based on the standard methods

(FRIEDENSTEIN et al., 1976; CAPLAN, 1991). Femurs were removed for bone

marrow cells acquirement by flushing BM cavities with DMEM containing low glucose

(Vitrocell Embriolife, Campinas, Brazil) supplemented with 10% fetal bovine serum

(Vitrocell Embriolife, Campinas, Brazil) and 0.1% penicillin (100 UI/mL) and

streptomycin (100 mg/mL) (Sigma Aldrich®, St. Louis, USA). Total bone marrow cells

were seeded in culture flasks and cultured in DMEM at 37 °C, 5% CO2 in a humidified

atmosphere. Every 3 days, medium was completely replenished and MSC growth and

morphology were monitored by bright field microscopy. When cells achieved 90%

confluence, they were split by trypsin method. MSC at passage 2 or 3 were used in

this study.

For characterization, MSC were stained with anti-CD271 (FITC, clone MLR2)

(Abcam, Cambridge, MA, USA) and evaluated by immunocytochemistry technique

(DOS SANTOS et al., 2017). Also, MSC were characterized by flow cytometry, as

described further, and the classic MSC multipotential differentiation capacities in

osteoblast and adipocyte were performed using a mouse mesenchymal stem cell

functional identification kit (SC010, R&D Systems, Abingdon, UK).

Mesenchymal-to-endothelial cell differentiation culture

Confluent MSC were washed with PBS and cultured for 15 days with endothelial

cell growth medium (EGM) (EGM-2®, Lonza, Walkersville, USA). Every 48 hours, the

medium was replenished and the cellular morphology and organization were monitored

42

by bright field microscopy. To evaluate the functionality of endothelial cells, 2 x 104

cells were seeded in a 96-well plate in 1:1 EGM and Matrigel® (Corning Inc.,

Tewskbury, USA). Cell cultures were observed by bright field microscopy every 24

hours for 15 days. As negative controls, MSC were seeded in 1:1 DMEM and Matrigel®

(Corning Inc., Tewskbury, USA).

Conditioned culture of bone marrow mononuclear or c-Kit+ cells and endothelial cell

supernatant

1 x 106 EC per mL were seeded with EGM medium (EGM-2®, Lonza,

Walkersville, USA) in 24-well culture plates and after 24 hours, the supernatant was

collected. 1 x 106 BM-MNC or c-Kit+ cells were seeded in 24-well culture plates with

1:1 McCoy 5A (Sigma Aldrich) supplemented with 10% fetal calf serum (Vitrocell

Embriolife, Campinas, Brazil), 0.1% penicillin and streptomycin (Sigma Aldrich, St.

Louis, USA), and supernatant from EC. After 72 hours, the non-adherent cells were

collected for cell cycle and immunophenotyping by flow cytometry or for RNA

extraction.

Co-culture of endothelial and c-Kit+ cells

1 x 106 EC per mL were seeded with 1:1 EGM medium (EGM-2®, Lonza,

Walkersville, USA) and McCoy 5A (Sigma Aldrich, St. Louis, USA) supplemented with

10% fetal calf serum (Vitrocell Embriolife, Campinas, Brazil) and 0.1% penicillin and

streptomycin (Sigma Aldrich, St. Louis, USA) in 24-well culture plates. Then, 5 x 105 c-

Kit+ cells were seeded on the EC and maintained in co-culture for 72 hours at 37 °C,

5% CO2 in a humidified atmosphere. After this period, the non-adherent cells were

collected for immunophenotyping by flow cytometry.

Conditioned culture of C1498 cells and endothelial cell supernatant


Walkersville, USA) in 24-well culture plates and after 24 hours, the supernatant was

collected. 2 x 105 C1498 cells (TIB-49ä, ATCCâ) were seeded in 24-well culture plates

with 1:1 DMEM containing low glucose (Vitrocell Embriolife, Campinas, Brazil)

supplemented with 10% fetal bovine serum (Vitrocell Embriolife, Campinas, Brazil),

0.1% penicillin (100 UI/mL) and streptomycin (100 mg/mL) (Sigma Aldrich®, St. Louis,

43

USA), and supernatant from endothelial cells. After 24 hours, cells were collected for

RNA extraction.

Cytokine quantification on endothelial cell supernatant


Walkersville, USA) in 24-well culture plates. After 24 hours, the supernatant was

collected and the concentrations of Ang-1, SCF, CXCL-12, IL-11, TGF-b, G-CSF, and

GM-CSF were determined by Enzyme Linked Immunosorbent Assay (ELISA) using

commercially available kits from R&D Systems (Quantikine ELISA®, R&D Systems,

Minneapolis, USA), except Ang-1 (Uscn Life Science Inc., Wuhan, China).

Flow cytometry

To access cell cycle, viability, apoptosis, and immunophenotype of

hematopoietic cells, ex vivo BM-MNC were collected. For cell cycle assay, cells were

fixed in 4% paraformaldehyde (Sigma Aldrich, St. Louis, USA), permeabilized with

0.1% Triton X-100 (Sigma Aldrich, St. Louis, USA), treated with RNase (BioRad,

Philadelphia, USA), and labeled with propidium iodide (PI) staining solution (BD

Pharmingen®, Becton Dickinson, New Jersey, USA). Once labeled, 1 x 104 cells were

acquired by flow cytometry. Cell cycle was assessed by quantifying the percentage of

histogram regions corresponding to G0/G1 and S/G2/M phases. For the viability and

apoptosis assays, cells were labeled with 8 μL of PI (BD Pharmingen®, Becton

Dickinson, New Jersey, USA) and 2.5 μL of annexin (BD Pharmingen®, Becton

Dickinson, New Jersey, USA). Once labeled, 1 x 104 cells were acquired by flow

cytometry. Viability analysis was performed by quantifying double-negative labeled

cells, and cells labeled with PI, annexin, or double-positive were considered apoptotic

cells. For hematopoietic cell immunophenotyping, cells (ex vivo bone marrow

mononuclear cells or post conditioned culture with endothelial supernatant or c-Kit+

cells post co-culture with endothelial cells) were labeled with antibody cocktails and a

viability stain (FVS780, BD Biosciences, New Jersey, USA). The antibodies used were

CD3-PE (145-2C11), CD11b-PE (M1/70), Ter119-PE (TER119), Ly6G-PE (RB6-8C5),

CD19-PE (MB19-1), CD16/32-PECy7 (2.4G2), CD34-FITC (RAM34), Thy1.1-PECy7

44

(OX-7), c-Kit-APC (2B8), Flk2-PE (A2F10.1), IL7r-FITC (SB/199), IL7r-PE (SB/199),

Sca-1-FITC (D7), Sca-1-PECy7 (D7), Sca-1-PE (D7), F4/80-APC (BM8), and CD11b-

FITC (M1/70), purchased from BD Biosciences. The populations evaluated were HSC

(Lin−Flk2−Thy1.1lowSca-1+c-Kit+), MPP (Lin−Flk2−Thy1.1lowSca-1−c-Kit+), CLP

(Lin−Il7rlowc-Kit+Sca-1+), CMP (Lin−Il7r−c-Kit+Sca-1−CD34+CD16/32low), GMP

(Lin−Il7r−c-Kit+Sca-1−CD34+CD16/32high), and MEP (Lin−Il7r−c-Kit+Sca-

1−CD34−CD16/32low). The flow cytometry strategy used is shown in Supporting

Information Figure 1 (Fig. S1a–e).

Endothelial and mesenchymal stem cells were stained with antibody cocktails

and a viability stain (FVS780, BD Biosciences). The antibodies used were CD90.1-PE-

Cy7 (OX-7), CD44-FITC (IM7), CD49e-PE (5H10-27), Sca-1-FITC (D7), CD105-APC

(266), CD34-APC (581), CD45-APC (30-F11), CD11b-FITC (M1-70), CD31-PE

(MEC13.3), CD144-PE (11D4.1), purchased from BD Biosciences (BD Pharmingen®,

Becton Dickinson, New Jersey, USA), and anti-CD133-PECy7 (315-2C11, BioLegend,

San Diego, USA).

Negative controls were performed by fluorescence minus one (FMO) strategy.

Data were acquired on a FACS Canto II (FACScan®, Becton Dickinson, New Jersey,

USA), and FlowJo® 10 software (Tree Star Inc., Ashland, USA) was used for data

analysis.

RNA isolation and quantitative real-time PCR

Total RNA was obtained from ex vivo and post co-culture bone marrow c-Kit+

cells, mesenchymal stem cells, endothelial cells, and post conditioned culture C1498

cells using a RNeasy RNA extraction kit (Qiagen, Germantown, MD) according to the

manufacturer’s protocol. Total RNA was reverse-transcribed into cDNA using the high-

capacity cDNA reverse transcription kit (Applied Biosystems, Foster City, CA).

Hematopoietic cells cDNA samples were amplified using the TaqMan universal

master mix with optimized concentrations of the primer set for Sox2

(Mm03053810_s1), Nanog (Mm02019550_s1), Pou5f1 (Mm03053917_g1), Gata1

(Mm02019550_s1), Gata2 (Mm02019550_s1), Gata3 (Mm02019550_s1), Sfpi1

(Mm02019550_s1), Ikzf3 (Mm02019550_s1), Nfe2 (Mm02019550_s1), Cebpa

(Mm02019550_s1), Il3ra (Mm00434273_m1), and Cxcr4 (Mm01292123_m1). Gene

45

expression was normalized to the housekeeping Gapdh (Mm99999915_g1) gene

expression.

Endothelial cells and/or mesenchymal stem cells cDNA samples were amplified

using the TaqMan universal master mix with optimized concentrations of the primer set

for Nt5e (Mm00501910_m1), Vcam1 (Mm01320970_m1), Mcam (Mm00522397_m1),

Flt1 (Mm00438980_m1), Kdr (Mm01222421_m1), Pdgfb (Mm00440677_m1), Nes

(Mm00450205_m1), Lepr (Mm00440181_m1) and Cspg4 (Mm00507257_m1). Gene

expression was normalized to the housekeeping Rn18s (Mm03928990_g1) gene

expression.

C1498 cells cDNA samples were amplified using the TaqMan universal master

mix with optimized concentrations of the primer set for Ccnd1 (Mm00432359_m1),

Ccne1 (Mm01266311_m1), Cdkn1a (Mm00432448_m1), Cdkn1b

(Mm00438168_m1). Gene expression was normalized to the housekeeping Gapdh

(Mm99999915_g1) gene expression.

The primer set was purchased from Applied Biosystems (Applied Biosystems,

Foster City, CA, USA). The gene expression was evaluated by real-time PCR using

StepOnePlus™ (Applied Biosystems, Foster City, CA) and quantified according to the

DDCt method (LIVAK e SCHMITTGEN, 2001).

Statistical analysis and reproducibility

Data sets passed through normality tests and were analyzed by Student’s t test

or analysis of variance (2way ANOVA) plus Tukey's post hoc test, unless otherwise

indicated. The level of significance adopted was 95% (p < 0.05) and all data are

represented as mean ± standard error of mean (SEM). n represents number of mice

analyzed in each experiment, as detailed in figure legends or tables. Statistical

analyses were performed using GraphPad Prism® 7 (GraphPad Software Inc., La Jolla,

USA) and the degrees of significance were indicated as follows: *p < 0.05, **p < 0.01,

***p < 0.001, and ****p < 0.0001.

Acknowledgments We acknowledge the assistance of the School of Pharmaceutical Science Flow

Cytometry and Animal Care Cores. This work was supported by Fundação de Amparo

46

à Pesquisa do Estado de São Paulo (FAPESP) grant (14/06872-2). R. A. Fock and P.

Borelli are fellows of the Conselho Nacional de Pesquisa e Tecnologia (CNPq).

Author contributions A. A. Hastreiter and R. A. Fock designed the project. A. A. Hastreiter performed

and analyzed all the experiments and wrote the manuscript. G. G. dos Santos provided

assistance for animal care and for blood samples collection. E. N. Makiyama and E.

W. C. Santos provided assistance for blood samples and flow citometry. R. A. Fock

supervised, helped to write the manuscript, reviewed and contributed to the drafting of

the manuscript.

COMPLIANCE WITH ETHICAL STANDARDS This study was approved by the Ethics Committee of the School of Pharmaceutical

Sciences at the University of São Paulo. The experiments comply with the current laws

of Brazil, where they were performed.

Conflicts of interests The authors declare that they have no conflict of interest.

47

Table 1. Effects of protein malnutrition on nutritional and hematopoietic parameters. Values for nutritional, peripheral blood, and bone marrow parameters are

expressed as mean ± standard error of mean. Asterisks indicate a significant difference

between groups: *(p≤0.05), **(p≤0.01), ***(p≤0.001), ****(p≤0.0001). n represents the

number of mice used in the experiments.

Variables Control Group

Malnourished Group

Nutritional parameters (n=20) (n=20) Food intake (g/day/animal) 3.84 ± 0.08 3.86 ± 0.07 Protein intake (g/day/animal) 0.508 ± 0.010 0.113 ± 0.002**** Body weight variation (%) 27.5 ± 1.1 -22.1 ± 1.2**** Total serum protein (g/dL) 5.63 ± 0.18 4.47 ± 0.09**** Serum albumin (g/dL) 1.90 ± 0.08 1.47 ± 0.06*** Peripheral blood parameters (n=20) (n=20) Erythrocytes (106/mm3) 8.83 ± 0.20 8.24 ± 0.24* Hemoglobin (g/dL) 12.74 ± 0.27 11.33 ± 0.34** Hematocrit (%) 38.95 ± 0.90 35.34 ± 0.96** Total leukocyte (/mm3) 2496.0 ± 23.6 957.1 ± 17.6**** Neutrophils (/mm3) 627.1 ± 1.4 244.4 ± 3.2*** Lymphocytes (/mm3) 1712.0 ± 148.5 694.3 ± 188.2** Monocytes (/mm3) 47.1 ± 11.8 11.8 ± 2.5 * Platelets (x103/mm3) 563.4 ± 41.6 612.8 ± 45.4 Myelogram (n=5) (n=5) Bone marrow cellularity (107/mm3) 4.06 ± 0.34 1.82 ± 0.19**** Blast cells (105/mm3) 12.93 ± 1.78 5.15 ± 0.67** Granulocyte precursors (105/mm3) 6.17 ± 0.88 2.42 ± 0.16** Band granulocytes (105/mm3) 22.24 ± 4.42 7.76 ± 0.75** Segmented granulocytes (105/mm3) 177.34 ± 14.39 87.63 ± 11.65*** Eosinophils (105/mm3) 11.17 ± 2.98 4.19 ± 0.88* Monocytes (105/mm3) 3.03 ± 0.73 0.71 ± 0.31** Lymphocytes (105/mm3) 52.97 ± 3.57 22.21 ± 2.32**** Pro-erythroblasts and basophilic erythroblasts (105/mm3)

16.15 ± 0.32 7.17 ± 0.62**

Polychromatophilic and orthochromatic erythroblasts (105/mm3)

102.01 ± 13.50 41.89 ± 4.89***

48

Fig. 1 Effects of protein malnutrition on hematopoietic stem and progenitor cells ex vivo. Percentage of G0/G1 (a) and S/G2/M (b) cell cycle phases. Representative cell

cycle histogram from control (c) and malnourished (d) mice. Percentage of viable (e)

and apoptotic cells (f). Percentage of HSC, MPP, CMP, GMP, and MEP (g).

Representative FACS plot of gate strategy of hematopoietic stem and progenitor cell

analyses of control (h) and malnourished (i) mice. Results referring to bone marrow

mononucleated cells in Control (n=5) and Malnourished (n=5) groups are expressed

as mean ± SEM. Heatmap of pluripotent, myeloid, and lymphoid differentiation gene

expression in c-Kit+ cells (j), values are relative to Gapdh expression. Significant

differences between groups are illustrated by *(p ≤ 0.05), **(p≤0.01) and ***(p ≤ 0.001).

n represents the number of animals used in the experiments.

49

Fig. 2 C1498 cells transplantation to control and malnourished mice. Design of

syngeneic transplantation (a). Quantification of C1498 cells in the bone marrow (b).

Representative FACS plot of C1498 quantification in control (c) and (d) malnourished

mice. Percentage of G0/G1, S, and G2/M cell cycle phases (e). Representative FACS

histogram of C1498 cell cycle in control (f) and malnourished (g) mice. Proliferation

index of bone marrow C1498 cells (h). Representative FACS proliferation histogram of

C1498 in control (i) and malnourished (j) mice. n=5 each group. Significant differences

between groups are illustrated by *(p ≤ 0.05). n represents the number of animals used

in the experiments.

50

Fig. 3 Characterization of endothelial cells differentiated from BM-MSC from control and malnourished mice. BM-MSC in vitro differentiation: osteoblasts stained

by May-Grünwald-Giemsa, optical magnitude 10x (a); osteoblasts stained by Alizarin

Red, optical magnitude 10x (b); osteoblasts stained positive for osteopontin, optical

magnitude 10x (c); adipocytes stained with oil red, optical magnitude 10x (d); and

adipocytes stained positive for FABP4, optical magnitude 40x (e). Heatmap of

immunophenotypic characterization of MSC and EC by flow cytometry (f), results are

expressed as log of positive cells. MSC morphology in bright field microscope: MSC

before induction of endothelial differentiation, optical magnitude 10x (g); MSC cultured

in Matrigel®, optical magnitude 10x (h); MSC after induction of endothelial

differentiation, optical magnitude 10x (i); and MSC after induction of endothelial

differentiation cultured in Matrigel®, optical magnitude 20x (j). Gene expression profile

for endothelial and mesenchymal characterization in Control and Malnourished groups:

Flt1 (k), Kdr (l), Vcam1 (m), Nt5e (n), Mcam (o), Pdgfb1 (p), Lepr (q), Nes (r) and Cspg4

(s). Results are relative to 18s expression and are expressed as mean ± SEM (n³8).

Different letters represent a statistically significant difference between the groups. n

represents the number of animals used in the experiments.

51

Fig. 4 Evaluation of the regulatory of CE to modulate hematopoiesis in PM.

Angpt1, Kitl, Cxcl12, Il11, Tgfb1, Igf1, Csf1, Csf2, and Csf3 gene expressions are

shown (a). Values are relative to 18s expression and are expressed as mean ± SEM

(n³8). Cytokine production by EC evaluation: Ang-1 (b), SCF (c), CXCL-12 (d), IL-11

(e), TGF-b (f) and G-CSF (g) are expressed as mean ± SEM (n=6). Significant

differences between groups are illustrated by *(p ≤ 0.05) and **(p≤0.01). n represents

the number of animals used in the experiments.

52

WITH CELL-CELL CONTACT a b c d e f

WITHOUT CELL-CELL CONTACT

g h i j k l

m n o

Control + Control EC

Control + Malnourished EC

Malnourished + Control EC

Malnourished + Malnourished EC

Fig. 5 Evaluation of the role of endothelial cells over hematopoietic differentiation. Evaluation of cell population after c-Kit+ cells and EC from control and

malnourished groups co-cultures in the quantification of: c-Kit+ cells (a), CMP (b), GMP

(c), MEP (d), Granulocytes (e), and Macrophages (f), n=3. Cell population after culture

of MNC and conditioned media with control EC supernatant or conditioned media with

malnourished EC supernatant in the quantification of: HSC (g), MPP (h), CLP (i), CMP

(j), GMP (k), and MEP (l), n=3. Gating strategy is described in Supplemental

Information (Fig. S1). Gene expression profile of c-Kit+ cells after conditioned media

with control EC supernatant and conditioned media with malnourished EC supernatant:

pluripotent genes (Sox2, Nanog, and Pou5f1) (m), myeloid differentiation (Gata1,

Gata2, Nfe2, Spi1, and Cebpa) (n), chemokine receptor (Il3ra and Cxcr4) (o), n=3,

values are relative to Gapdh expression. Significant differences between groups are

illustrated by *(p ≤ 0.05), **(p≤0.01) and ***(p ≤ 0.001). Different letters represent a

statistically significant difference between the groups. n represents the number of

animals used in the experiments.

Control Malnourished0

20

40

60

80

c-K

it+ cel

ls (%

of c

ells

)

c-Kit+ cells

a abb ab

Control Malnourished0.0

0.2

0.4

0.6

0.8

CM

P (%

of c

ells

)

CMP

a a

bb


1

2

3

4

GM

P (%

of c

ells

)

GMP

a

b

c

a


2

4

6

8

ME

P (%

of c

ells

)

MEP

aba

b

a


10

20

30

40

Gra

nulo

cyte

s (%

of c

ells

)

Granulocytes

a

bb

c


0.02

0.04

0.06

0.08

Mac

roph

ages

(% o

f cel

ls)

Macrophages


0.2

0.4

0.6

HSC

(% o

f cel

ls)

ab

c

d

HSC


0.2

0.4

0.6

0.8

MPP

(% o

f cel

ls)

a

a

bbMPP


0.01

0.02

0.03

0.04

CLP

(% o

f cel

ls)

CLP

a

b b b


0.1

0.2

0.3

0.4

0.5

CM

P (%

of c

ells

)

CMP

a a

b

a


0.5

1.0

1.5

2.0

GM

P (%

of c

ells

)

GMP

aa

b

a


0.05

0.10

0.15

0.20

0.25

MEP

(% o

f cel

ls)

MEP

a a

b ab

Sox2 Nanog Pou5f10.0

0.5

1.0

1.5

2.0

mR

NA

(re

lati

ve t

o Gapdh

)

***

*** *

Gata1 Gata2 Nfe2 Sfpi1 Cebpa0

2

4

68

10

mRN

A (r

elat

ive

to G

apdh

)

*****

*****

**

*

Il3ra Cxcr40

2

4

6

mR

NA

(re

lati

ve t

o G

apd

h)

*** Cond. Control EC

Cond. Malnourished EC

Cond. Control EC

Cond. Malnourished EC

53

Fig. 6 Effects of conditioned media with control and malnourished EC supernatant over viability and cell cycle. BM-MNC viability (a) and apoptosis status (b) after culture with culture

media alone and culture media conditioned with control or malnourished EC supernatant.

Percentage of BM-MNC G0/G1 (c) and S/G2/M (d) cell cycle phases after culture media

conditioned with control or malnourished EC supernatant, n=3 each group. Representative

FACS histogram of cell cycle of control BM-MNC conditioned with control EC supernatant (e),

malnourished BM-MNC conditioned with control EC supernatant (f), control BM-MNC

conditioned with malnourished EC supernatant (g), and malnourished BM-MNC conditioned

with malnourished EC supernatant (h). Results of Ccnd1 (i), Ccne1 (j), Cdkn1b (k), and

Cdkn1a (l) gene expression on C1498 cells after conditioned culture with control or

malnourished EC supernatant. Values are relative to Gapdh expression and are expressed as

mean ± SEM (n=5). Significant differences between groups are illustrated by *(p ≤ 0.05)

and ***(p ≤ 0.001); n represents the number of animals used in the experiments.

54

SUPPLEMENTARY MATERIAL

Fig. S1 Flow cytometry gates strategy. Forward Scatter x Side Scatter, single cells and

viable cells gates (a). Hematopoietic stem cells (R2) and hematopoietic multipotent progenitors

(R3) gates strategy (b). Common lymphoid progenitors (R2) gates strategy (c). Common

myeloid progenitors (R2), granule-monocytic progenitors (R3), and megakaryocytic-erythroid

progenitors (R4) gates strategy (d). Granulocytes (R2) and macrophages (R3) gates strategy

(e).

55

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SASINE, J. P.; YEO, K. T.; CHUTE, J. P. Concise Review: Paracrine Functions of Vascular Niche Cells in Regulating Hematopoietic Stem Cell Fate. Stem Cells Transl Med, v. 6, n. 2, p. 482-489, Feb 2017. ISSN 2157-6564 (Print) 2157-6564 (Linking). Disponível em: < https://www.ncbi.nlm.nih.gov/pubmed/28191767 >. SCHOFIELD, R. The relationship between the spleen colony-forming cell and the haemopoietic stem cell. Blood Cells, v. 4, n. 1-2, p. 7-25, 1978. ISSN 0340-4684 (Print) 0340-4684 (Linking). Disponível em: < http://www.ncbi.nlm.nih.gov/pubmed/747780 >. SEMERAD, C. L. et al. A role for G-CSF receptor signaling in the regulation of hematopoietic cell function but not lineage commitment or differentiation. Immunity, v. 11, n. 2, p. 153-61, Aug 1999. ISSN 1074-7613 (Print) 1074-7613 (Linking). Disponível em: < https://www.ncbi.nlm.nih.gov/pubmed/10485650 >. VITURI, C. L. et al. Alterations in proteins of bone marrow extracellular matrix in undernourished mice. Braz J Med Biol Res, v. 33, n. 8, p. 889-95, Aug 2000. ISSN 0100-879X (Print) 0100-879X (Linking). Disponível em: < http://www.ncbi.nlm.nih.gov/pubmed/10920430 >. WADHWA, M.; THORPE, R. Haematopoietic growth factors and their therapeutic use. Thromb Haemost, v. 99, n. 5, p. 863-73, May 2008. ISSN 0340-6245 (Print) 0340-6245 (Linking). Disponível em: < http://www.ncbi.nlm.nih.gov/pubmed/18449415 >. WEISSMAN, I. L.; SHIZURU, J. A. The origins of the identification and isolation of hematopoietic stem cells, and their capability to induce donor-specific transplantation tolerance and treat autoimmune diseases. Blood, v. 112, n. 9, p. 3543-53, Nov 1 2008. ISSN 1528-0020 (Electronic) 0006-4971 (Linking). Disponível em: < http://www.ncbi.nlm.nih.gov/pubmed/18948588 >. XAVIER, J. G. et al. Protein-energy malnutrition alters histological and ultrastructural characteristics of the bone marrow and decreases haematopoiesis in adult mice. Histol Histopathol, v. 22, n. 6, p. 651-60, Jun 2007. ISSN 1699-5848 (Electronic) 0213-3911 (Linking). Disponível em: < https://www.ncbi.nlm.nih.gov/pubmed/17357095 >. XU, C. et al. Stem cell factor is selectively secreted by arterial endothelial cells in bone marrow. Nat Commun, v. 9, n. 1, p. 2449, Jun 22 2018. ISSN 2041-1723 (Electronic) 2041-1723 (Linking). Disponível em: < https://www.ncbi.nlm.nih.gov/pubmed/29934585 >.

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4 CAPÍTULO II

EFEITOS DA DESNUTRIÇÃO PROTEICA SOBRE ASPECTOS REGULATÓRIOS DA HEMATOPOESE DAS CÉLULAS TRONCO MESENQUIMAIS MEDULARES

63

Title: Effects of protein malnutrition on hematopoietic regulatory aspects of

bone marrow mesenchymal stem cells

Authors: Araceli Aparecida Hastreiter1, Guilherme G. dos Santos1, Edson Naoto

Makiyama1, Ed Wilson Cavalcante Santos1, Primavera Borelli1, Ricardo Ambrósio

Fock1*


Sciences, University of São Paulo, São Paulo, Bra





[email protected]

64

ABSTRACT Protein malnutrition (PM) causes anemia and leukopenia as it reduces hematopoietic

precursors and impairs the production of mediators that regulate hematopoiesis.

Hematopoiesis occurs in distinct bone marrow (BM) niches, which modulate the

processes of differentiation, proliferation and self-renewal of hematopoietic stem cells

(HSC). Mesenchymal stem cells (MSC) contribute to biochemical composition of BM

niches by the secretion of several growth factors and cytokines and play an important

role in the regulation of HSC and hematopoietic progenitors. In this study, we

investigated the effect of PM on the hematopoietic regulatory function of MSC.

C57BL/6 mice were divided into control and malnourished groups, which received,

respectively, a normal protein diet (12% casein) and a low protein diet (2% casein).

PM altered the synthesis of CXCL-12, SCF, Ang-1 and TFG-β by MSC, indicating that

malnourished MSC are in a pro-proliferative status. However, hematopoietic cells from

malnourished group did not respond to MSC stimuli as control group. In addition,

malnourished MSC affected the hematopoietic differentiation capacity, decreasing the

lymphoid, granulocytic and megakaryocytic-erythroid differentiation of in control group.

Nevertheless, malnourished MSC only downregulated the megakaryocytic-erythroid

differentiation in malnourished group. Therefore, we infer hematopoietic alterations

caused by PM are due multifactorial alterations and, at least in part, MSC contribute to

hematological impairment.

Key-words: Protein malnutrition; Mesenchymal stem cell; Hematopoiesis regulation.

65

1. INTRODUCTION

Malnutrition is a multifactorial nutritional disorder that affects more than 10% of

the global population and is reflected in metabolic alterations dependent on the degree

and duration of malnutrition, even as the presence of comorbidities (NORMAN et al.,

2011; FAO, 2018). Protein malnutrition (PM) is the most common type of malnutrition

and mainly affects patients with chronic diseases, children and the elderly (PEDRUZZI

e TEIXEIRA, 2007). PM can disrupt all tissues, especially those with high cellular

turnover, such as hematopoietic tissue (BORELLI et al., 2004; XAVIER et al., 2007;

BORELLI et al., 2009; SANTOS et al., 2017).

PM causes alterations in hematopoietic organs leading to anemia, leukopenia

and impairment of the immune response (KEUSCH, 1994; BORELLI et al., 1995;

VITURI et al., 2000; KEUSCH, 2003; BORELLI et al., 2007; FOCK et al., 2007;

SANTOS et al., 2017). Additionally, PM induces cell cycle arrest in hematopoietic

progenitors and can yield to bone marrow (BM) hypoplasia (BORELLI et al., 2009;

NAKAJIMA et al., 2014).

The regulation of self-renewal and differentiation of hematopoietic stem cell

(HSC) and progenitors – multipotent progenitors (MPP), common lymphoid progenitors

(CLP), common myeloid progenitors (CMP), granule-monocytic progenitors (GMP) and

megakaryocytic-erythroid progenitors (MEP) – is exercised by the BM niche cells,

through direct and indirect mechanisms, such as cytokines and growth factors

synthesis, as well as cell-cell contact (KIEL et al., 2005; WEISSMAN e SHIZURU,

2008).

Mesenchymal stem cells (MSC) are cells that play an important role in the BM

niche formation as well as in the modulation of hematopoiesis. In spite of MSC may

modulate hematopoietic differentiation and maturation, their most important role is the

regulation of HSC and hematopoietic progenitors (GARCIA-GARCIA et al., 2015).

MSC expressing leptin receptor (LepR+) have been shown to induce HSC quiescence

by release of Ang-1 (ZHOU et al., 2015), whereas Nestin+ (Nes+) MSC and "CAR cells"

– reticular cells that synthesize large amounts of CXCL-12 – regulate HSC

maintenance through SCF and CXCL-12 synthesis (SUGIYAMA et al., 2006;

MENDEZ-FERRER et al., 2010; EHNINGER e TRUMPP, 2011; DING et al., 2012). In

addition, MSC produce TGF-β, a negative regulator of hematopoiesis, which inhibits

the cell cycle of the hematopoietic progenitors (RUSCETTI et al., 2005).

66

In this study, we show that PM increased pluripotency and induced a pro-

proliferative profile in MSC in vitro, which resulted in an increase in HSC, MPP and

CMP, as well as in the decrease of the lineage-specific differentiation. However,

malnourished MSC failed to improve proliferation in malnourished animals and, in

addition, downregulated megakaryocytic-erythroid differentiation.

2. MATERIALS AND METHODS Mice and diets

This study was approved by the Ethics Committee of the School of

Pharmaceutical Sciences at the University of São Paulo. Male mice of the C57BL/6 inbred strains of 45-60-days-old were maintained in individual cages at 71 ± 37 °F, and

relative humidity at 55% ± 10%, with a regular 12-hour light/dark cycle. Mice underwent

an adaptation period (10 to 15 days), in which all animals received normoproteic diet

and water ad libitum until stabilization of body weight. After this period, mice were

divided into two groups, which received either normoproteic diet (control group) or

hypoproteic diet (malnourished group).

Normoproteic and hypoproteic diets were prepared inhouse. Mineral and

vitamin mixtures were prepared according to the recommendations of the American

Institute of Nutrition (AIN-93M) for adult mice (REEVES et al., 1993; REEVES, 1997).

The protein source used was casein (> 85% protein) and normoproteic and hypoproteic

diets contained 12% and 2%, respectively. Both diets contained 100 g kg-1 sucrose, 80

g kg-1 soybean oil, 10 g kg-1 fiber, 2.5 gk g-1 choline bitartrate, 1.5 g kg-1 L-methionine,

40 g kg-1 of mineral mixture and 10 g kg-1 of vitamin mixture. The control diet contained

120 g kg-1 casein and 636 g kg-1 cornstarch, while the malnourishment diet contained

20 g kg-1 casein and 736 g kg-1 cornstarch. With the exception of the protein and corn

starch content, the two diets were identical and isocaloric, providing 1716.3 kJ/100 g.

The final protein content of both diets was confirmed by the standard micro-Kjeldahl

method.

The period for the induction of malnutrition was 35 to 40 days, and the

nutritional evaluation was performed by monitoring body weight, food consumption and

protein intake every 48 hours during the experimental period (XAVIER et al., 2007;

DOS SANTOS et al., 2017). The variation in body weight was calculated as a relative

67

value between the body weight on the first day of induction to malnutrition and the last

day of this period.

Hemogram

Blood samples were collected with EDTA (Merck, Darmstadt, Germany) from

both control and malnourished animals. Hemograms were obtained by loading blood

samples into ABX Micros ABC Vet® equipment (Horiba ABX, Montpellier, France). The

morphological and leukocyte differential analyses were performed on blood smears

stained by May-Grünwald-Giemsa (Merck, Darmstadt, Germany) technique.

Serum protein and albumin quantification

After malnourishment induction, mice were euthanized and blood samples were

collected serum was separated by centrifugation (1000 x g for 10 minutes at 4°C). The

concentrations of serum proteins and albumin were determined by the use of

commercial kits (Labtest Diagnóstica SA, Lagoa Santa, Brazil) and based on standard

methods.

Myelogram

Bone marrow cells were obtained by flushing femurs with Dulbecco’s modified

Eagle’s medium containing low glucose (DMEM) (Vitrocell Embriolife, Campinas,

Brazil) supplemented with 10% fetal calf serum (Vitrocell Embriolife, Campinas, Brazil),

0,1% Penicillin and Streptomycin (Sigma Aldrich, St. Louis, USA). Bone marrow

cellularity was determined by counting obtained cells using a Neubauer

hemocytometer and myelogram was performed by morphological and differential

analysis on cytocentrifugated smears stained by May-Grünwald-Giemsa standard

method.

Mesenchymal stem cells isolation and characterization

MSC were obtained and characterized based on the standard methods

(FRIEDENSTEIN et al., 1976; CAPLAN, 1991). Femurs were removed for bone

68

marrow cells acquirement by flushing BM cavities with Dulbecco’s modified Eagles

medium (DMEM) containing low glucose (Vitrocell Embriolife, Campinas, Brazil)

supplemented with 10% fetal bovine serum (Vitrocell Embriolife, Campinas, Brazil) and

0,1% penicillin (100 UI/mL) streptomycin (100 mg/mL) (Sigma Aldrich®, St. Louis,

USA). Total bone marrow cells were seeded in culture flasks and cultured in DMEM at

98.6oF 5% CO2 in a humidified atmosphere. Every 3 days, medium was completely

replenished and MSC growth and morphology were monitored by bright field

microscopy. When cells achieved 90% confluence, they were splited by trypsin

method. MSC at passage 2 or 3 were used in this study.

For characterization, MSC were characterized by flow cytometry. MSC were

stained with antibody cocktails and a viability stain (FVS780, BD Biosciences). The

antibodies used were CD90.1-PE-Cy7 (OX-7), CD44-FITC (IM7), CD49e-PE (5H10-

27), CD34-APC (581), CD45-APC (30-F11), CD11b-FITC (M1-70), purchased from BD

Biosciences (BD Pharmingen®, Becton Dickinson, New Jersey, USA). To stablish negative controls, we performed unstained and stained cells with fluorescence-minus-

one (FMO) control stain sets. Data were acquired on a FACS Canto II (FACScan®,

Becton Dickinson, New Jersey, USA) and FlowJo® 10 software (Tree Star Inc,

Ashland, USA) was used for data analysis.

In addition, MSC were stained with anti-CD271 (FITC, clone MLR2) (Abcam,

Cambridge, MA, USA) and evaluated by immunocytochemistry technique and the

classic MSC multipotential differentiations capacity in osteoblast and adipocyte were

performed using a mouse mesenchymal stem cell functional identification kit (SC010,

R&D Systems, Abingdon, UK), as described previously (DOS SANTOS et al., 2017).

Cytokine quantification on mesenchymal stem cells supernatant

1x106 mesenchymal stem cells per mL were seeded with DMEM containing low

glucose (Vitrocell Embriolife, Campinas, Brazil) supplemented with 10% fetal bovine

serum (Vitrocell Embriolife, Campinas, Brazil) and 0,1% penicillin (100 UI/mL)

streptomycin (100 mg/mL) (Sigma Aldrich®, St. Louis, USA) in 24 wells culture plates.

After 24 hours, the supernatant was collected and the concentrations of Ang-1, SCF,

CXCL-12, IL-11, IL-3, TGF-b, G-CSF and GM-CSF were determined by Enzyme

Linked Immuno Sorbent Assay (ELISA) using commercially available kits from R&D

69

Systems (Quantikine ELISA®, R&D Systems, Minneapolis, USA), except Ang-1 (Uscn

Life Science Inc., Wuhan, China).

Bone marrow mononuclear and c-Kit+ cells isolation

Total bone marrow cells were flushed with McCoy 5A (Sigma Aldrich, St. Louis,

USA) supplemented with 10% fetal calf serum (Vitrocell Embriolife, Campinas, Brazil),

0,1% Penicillin and Streptomycin (Sigma Aldrich, St. Louis, USA) of both femurs and

tibias, then mononuclear cells were separated by density gradient with Ficoll-

Histopaque technique (Sigma Aldrich, St. Louis, USA). After that, c-Kit+ were

separated using magnetic-activated cell sorting (MACS). First, mononuclear cells were

labeled with anti-CD117 microbeads (Miltenyi Biotech Inc., Auburn, EUA) and c-Kit+

cells were isolated on a magnetic column following the manufacturer´s instructions.

Conditioned culture of bone marrow mononuclear or c-Kit+ cells and mesenchymal

stem cells supernatant

1x106 mesenchymal stem cells per mL were seeded with DMEM containing

low glucose (Vitrocell Embriolife, Campinas, Brazil) supplemented with 10% fetal

bovine serum (Vitrocell Embriolife, Campinas, Brazil) and 0,1% penicillin (100 UI/mL)

streptomycin (100 mg/mL) (Sigma Aldrich®, St. Louis, USA) in 24 wells culture plates

and after 24 hours, the supernatant was collected. 1x106 bone marrow mononuclear

cells or c-Kit+ cells were seeded in 24 wells culture plates with 1:1 McCoy 5A (Sigma

Aldrich) supplemented with 10% fetal calf serum (Vitrocell Embriolife, Campinas,

Brazil), 0,1% Penicillin and Streptomycin (Sigma Aldrich, St. Louis, USA) and

supernatant from mesenchymal stem cells. After 72 hours, the non-adherent cells were

collected for cell cycle and immunophenotyping by flow cytometry or for RNA

extraction, as described above.

Co-culture of mesenchymal stem and c-Kit+ cells

1x106 mesenchymal stem cells per mL were seeded 1:1 with DMEM containing

low glucose (Vitrocell Embriolife, Campinas, Brazil) supplemented with 10% fetal

70

bovine serum (Vitrocell Embriolife, Campinas, Brazil) and 0,1% penicillin (100 UI/mL)

streptomycin (100 mg/mL) (Sigma Aldrich®, St. Louis, USA) and McCoy 5A (Sigma

Aldrich, St. Louis, USA) supplemented with 10% fetal calf serum (Vitrocell Embriolife,

Campinas, Brazil), 0,1% Penicillin and Streptomycin (Sigma Aldrich, St. Louis, USA)

in 24 wells culture plates. Then, 5x105 c-Kit+ cells were seeded on the mesenchymal

stem cells and maintained in co-culture for 72 hours at 98.6oF 5% CO2 in a humidified

atmosphere. After this period, the non-adherent cells were collected for

immunophenotyping by flow cytometry, as described above.

Flow cytometry of hematopoietic cells

To access cell cycle, viability, apoptosis and immunophenotyping of

hematopoietic cells, bone marrow mononuclear cells post conditioned culture with

mesenchymal stem cells supernatant or c-Kit+ cells post co-culture with mesenchymal

stem cells were collected. For cell cycle assay, cells were fixed in 4%

paraformaldehyde (Sigma Aldrich, St. Louis, USA), permeabilized with 0.1% of Triton

X-100 (Sigma Aldrich, St. Louis, USA), treated with RNase (BioRad, Philadelphia,

USA) and labeled with Propidium Iodide Staining Solution (BD Pharmingen®, Becton

Dickinson, New Jersey, USA). Once labeled, 1x104 cells were acquired by flow

cytometry. Cell cycle was assessed by quantifying the percentage of histogram regions

corresponding to G0/G1 and S/G2/M. For the viability and apoptosis assay, cells were

labeled with 8 μl of PI (BD Pharmingen®, Becton Dickinson, New Jersey, USA) and

2.5 μl of annexin (BD Pharmingen®, Becton Dickinson, New Jersey, USA). Once

labeled, 1x104 cells were acquired by flow cytometry. Viability analysis was performed

by quantifying double-negative labeled cells and cells labeled with PI, annexin or

double-positive were considered apoptotic cells. For hematopoietic cells immunophenotyping, cells (bone marrow mononuclear

cells post conditioned culture with mesenchymal stem cells supernatant or c-Kit+ cells

post co-culture with mesenchymal stem cells) were labeled with antibody cocktails and

a viability stain (FVS780, BD Biosciences, New Jersey, USA). The antibodies used

were CD3-PE (145-2C11), CD11b-PE (M1/70), Ter119-PE (TER119), Ly6G-PE (RB6-

8C5), CD19-PE (MB19-1), CD16/32-PECy7 (2.4G2), CD34-FITC (RAM34), Thy1.1-

PECy7 (OX-7), c-Kit-APC (2B8), Flk2-PE (A2F10.1), IL7r-FITC (SB/199), IL7r-PE

71

(SB/199), Sca-1-FITC (D7), Sca-1-PECy7 (D7), Sca-1-PE (D7), F4/80-APC (BM8) and

CD11b-FITC (M1/70), purchased from BD Biosciences. The populations evaluated

were hematopoietic stem cells (HSC - Lin-Flk2-Thy1.1lowSca-1+c-Kit+), hematopoietic

multipotent progenitors (MPP - Lin-Flk2-Thy1.1lowSca-1-c-Kit+), comum lymphoid

progenitors (CLP - Lin-Il7rlowc-Kit+Sca-1+), comum myeloid progenitors (CMP - Lin-Il7r-

c-Kit+Sca-1-CD34+CD16/32low), granule-monocytic progenitors (GMP - Lin-Il7r-c-

Kit+Sca-1-CD34+CD16/32high) and megakaryocytic-erythroid progenitors (MEP - Lin-

Il7r-c-Kit+Sca-1-CD34-CD16/32low). The flow cytometry strategy used is shown in

Supporting Information Figure 1.

To stablish negative controls, we performed unstained and stained cells with

fluorescence-minus-one (FMO) control stain sets. Data were acquired on a FACS

Canto II (FACScan®, Becton Dickinson, New Jersey, USA) and FlowJo® 10 software

(Tree Star Inc, Ashland, USA) was used for data analysis.

RNA isolation and quantitative real-time PCR

Total RNA was obtained from mesenchymal stem cells and from post

conditioned culture bone marrow c-Kit+ cells using a RNeasy RNA extraction kit

(Qiagen, Germantown, MD) according to the manufacturer’s protocol. Total RNA was

reverse-transcribed into cDNA using the High-capacity cDNA reverse transcription kit

(Applied Biosystems, Foster City, CA).

cDNA samples from MSC were amplified in the TaqMan universal master mix

with optimized concentrations of the primer set for Angpt1 (Mm00456503_m1), Kitl

(Mm00442972_m1), Cxcl12 (Mm00445553_m1), Prom1 (Mm00477115_m1), Il11

(Mm00434162_m1), Il3 (Mm00439631_m1), Tgfb1 (Mm01178820_m1), Igf1

(Mm00439560_m1), Csf1 (Mm00432686_m1), Csf2 (Mm01290062_m1), Csf3

(Mm00438335_g1), Wnt3a (Mm00437337_m1), Wnt5a (Mm00437347_m1), Icam1

(Mm00516023_m1), Eng (Mm00468256_m1), Mcam (Mm00522397_m1), Pdgfb

(Mm00440677_m1), Nes (Mm00450205_m1), Lepr (Mm00440181_m1) and Cspg4

(Mm00507257_m1). Gene expression was normalized to the housekeeping Rn18s

(Mm03928990_g1) gene expression.

cDNA samples from hematopoietic cells were amplified in the TaqMan universal

master mix with optimized concentrations of the primer set for Sox2

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(Mm03053810_s1), Nanog (Mm02019550_s1), Pou5f1 (Mm03053917_g1), Gata1

(Mm02019550_s1), Gata2 (Mm02019550_s1), Gata3 (Mm02019550_s1), Sfpi1

(Mm02019550_s1), Ikzf3 (Mm02019550_s1), Nfe2 (Mm02019550_s1), Cebpa

(Mm02019550_s1), Il3ra (Mm00434273_m1) and Cxcr4 (Mm01292123_m1) Gene

expression was normalized to the housekeeping Gapdh (Mm99999915_g1) gene

expression.

The primer set was purchased from Applied Biosystems (Applied Biosystems,

Foster City, CA, USA). The gene expression was evaluated by real-time PCR using

StepOnePlusTM (Applied Biosystems, Foster City, CA) and quantified according to the

DDCt method (LIVAK e SCHMITTGEN, 2001).

Statistical analysis and reproducibility

Data sets passed through normality tests and were analyzed by Student’s t test

or analysis of variance (2way ANOVA) plus Tukey's post hoc test. All data are

represented as mean ± SEM, unless otherwise indicated. The level of significance

adopted was 95% (p < 0.05) and n represents number of mice analyzed in each

experiment, as detailed in figure legends or tables. Statistical analyzes were performed

using GraphPad Prism® 8 (GraphPad Software Inc., La Jolla, USA). *p < 0.05, **p <

0.01, ***p < 0.001, ****p < 0.0001.

3. RESULTS

Nutritional status

In this study, we used a low-protein diet to induce protein malnutrition. Mice from

both groups exhibited a similar food intake (Fig. 1A) during the period of malnutrition

induction, however the malnourished group had a lower protein intake (Fig. 1B) due to

hypoproteic diet. As consequence, malnourished mice presented body weight loss and

decrease of total serum protein and albumin concentrations (Fig. 1C-E).

Hematological evaluation

73

The hematologic evaluation was performed in mice from control and

malnourished mice by hemogram and myelogram, described in Table 1. Mice that

received hypoproteic diet (PM group) exhibited expressive leukopenia, with decreased

number of neutrophils, lymphocytes and monocytes, but cellular morphological

differences between groups were not found. PM group also showed a hypoplasic bone

marrow and a significant reduction in the total nucleated cell count and in the absolute

value of all lineages. In addition, PM group showed a decreased quantification of

erythrocytes, concentration of hemoglobin and hematocrit values, characteristic of PM.

Mesenchymal stem cell characterization

In order to evaluate the impact of PM on MSC, we first collected and isolated

BM-MSC. MSC were immunophenotypically characterized by flow cytometry technique

and the cells from both groups stained positively for the mesenchymal surface markers

CD44, CD49e and CD90.1, whereas did not stain for the hematopoietic markers CD34,

CD45 and CD11b, and no significant differences were observed between control and

malnourished groups. The osteoblastic and adipocytic differentiation capacity of MSCs

from control and malnourished animals were confirmed and no differences between

groups were observed (data not shown).

In addition, we evaluated the mRNA expression from function-related and

widely expressed genes in MSC. PM suppressed the expression of Igf1 in MSC, but

did not affect the expression of Icam1, Pdgf1, Eng, Mcam and Prom1 (Fig. 2B). Since MSC are not a homogeneous cell population, we performed the gene expression of nestin

(Nes), NG2 (Cspg4) and leptin receptor (Lepr), in order to evaluate if PM could alter the population subtype of MSC. MSC from malnourished group exhibited decreased expression of Nes and Cspg4, but

no differences were observed on Lepr expression (Fig. 2B).

Protein malnutrition increases pro-proliferative status of MSC

To evaluate the hematopoietic modulatory properties of MSC, the production of

hematopoietic regulatory cytokines SCF, TGF-b, Ang-1, CXCL-12, IL-11, IL-3, GM-

CSF and G-CSF was quantified on the supernatant of MSC cultures from both groups.

Additionally, the expression of genes related to HSC maintenance and hematopoietic

progenitor/precursor differentiation was also evaluated by qPCR.

74

MSC from malnourished group produced higher amount of SCF (Fig. 3A) and

its respective gene (Kitl) expression was upregulated (Fig. 3B) in comparison to MSC

from the control group. On the other hand, Ang-1 and Angpt1 were downregulated in

malnourished group (Fig. 3C and Fig. 3F). TGF-b and CXCL-12 quantifications were

decreased in malnourished group (Fig. 3B and Fig. 3D), even though there were no

differences in Tgfb1 or Cxcl12 gene expression between groups (Fig. 3F).

Concerning the regulation of the hematopoietic differentiation, no alterations on

G-CSF protein and mRNA (Csf3) and on GM-CSF and M-CSF encoding gene (Csf2

and Csf1, respectively) expression levels were observed between groups (Fig. 4E-F).GM-CSF and IL-11 protein levels were not detected, although we observed a

significative increase in Il11 mRNA expression (Fig. 3F). IL-3 was not detected in both

control and PM groups, neither by ELISA nor Il3 gene expression quantification by

qPCR.

MSC alters pluripotency gene expression and suppresses differentiation-related genes

expression in protein malnutrition

Since PM affected the function of MSC, we performed a gene expression profile

in order to evaluate the effect of MSC from control and malnourished groups on c-Kit+

cells after conditionate cultures with MSC supernatant. The expression of the

pluripotency genes Sox2, Pou5f1 (Oct-4) and Nanog did not show difference in c-Kit+

cells from control and malnourished groups when cultivated with their respective

conditioned medium (Fig. 4A). However, c-Kit+ cells from control group cultivated with

conditioned medium from malnourished group showed that the gene expression of

Sox2, Pou5f1 (Oct-4) and Nanog were upregulated in comparison to the other groups

studied (Fig. 4A) Additionally, c-Kit+ cells from malnourished group when cultured with

malnourished conditioned medium showed increased expression for Sox2 in

comparison to c-Kit+ cells from malnourished group cultured with control conditioned

medium (Fig. 4A).

PM did not affect the expression of Il3ra (IL-3 receptor) and Cxcr4 (CXCL-12

receptor) in all groups studied (Fig. 4B), but conditioned medium from malnourished

groups downregulated the expression of the lymphoid (Ikzf3 and Gata3, Fig. 4C) and

myeloid (Gata1, Gata2, Sfpi1 and Cebpa, Fig. 4D) differentiation-related genes in c-

75

Kit+ cells, but none differences were observed in the gene expression of Nfe2 (Fig. 4C).

Effects of MSC on viability and cell cycle modulation in hematopoietic cells in PM

Since malnourished MSC produced increased amount of SCF, a pro-

proliferative growth factor, and decreased Ang-1 and CXCL-12 production, which

induce HSC quiescence and prevent apoptosis, the effect of MSC supernatants on the

viability and cell cycle of hematopoietic cells was investigated. BM-MNC from both

groups were cultured with MSC supernatant from control or malnourished group and

the viability, apoptosis status and cell cycle were evaluated by flow cytometry. No

differences were found between groups in the viability on the cultures performed with

control or malnourished conditioned media (Fig. 5A), although the viability of cultures

was increased when compared to culture media alone (data not shown).

However, the conditioned cultures with MSC supernatant induced quantitative

alterations in cell cycle phases. Cells from control group were more frequent in S/G2/M

cell cycle phases when cultured with malnourished MSC conditioned media and,

consequently, less frequent in G0/G1 cell cycle phases, but there was no difference in

the cell cycle phases in cells from malnourished group after control or malnourished

conditionate culture (Fig. 5B).

Protein malnutrition modulates the paracrine effects of MSC on the proliferation and

differentiation of hematopoietic cells

In order to investigate the ability of MSC to induce hematopoietic differentiation,

we first performed BM-MNC cultures conditioned with MSC supernatant of both

groups. Malnourished conditioned medium increased HSC, MPP and CMP populations

in control group, whereas decreased CLP, GMP and MEP populations (Fig. 5C and

Fig. 5D). However, malnourished MSC supernatant did not impact on alterations in the

quantification of hematopoietic stem and progenitor cells in malnourished group,

except in MEP quantification, which was reduced by malnourished MSC supernatant

(Fig. 5D).

Effect of MSC – c-Kit+ cells contact on lineage-specific differentiation in PM

76

Since malnourished MSC paracrine signaling altered the populations of

hematopoietic progenitors, we inquired if the cell-cell contact would intensify or not the

c-Kit+ cells differentiation into lineage-specific cells. MSC from both groups were cross-

cultured with c-Kit+ cells, and the differentiation line from CMP to granulocytic cells was

evaluated. Interestingly, the cultures performed with MSC-c-Kit+ cells contact were

able to maintain cell viability (Fig. 6A).

Malnourished MSC decreased c-Kit+ cells, GMP, granulocytes and

monocytes/macrophages in both groups, but not statistically different (Fig. 6B-D).

Malnourished group exhibited fewer CMP and MEP cells when compared to control

group after co-culture with control MSC, but no differences were found in CMP and

MEP quantification between groups after co-culture with malnourished MSC (Fig. 6C).

4. DISCUSSION

Protein malnutrition (PM) causes anemia and leukopenia as it reduces

hematopoietic stem and progenitor cells and impairs the production of mediators that

induce hematopoiesis (BORELLI et al., 2007; XAVIER et al., 2007; SANTOS et al.,

2017). HSC provides all mature hematopoietic cells, through a controlled balance

between self-renewal and differentiation, which mainly depends on the cells from BM

niche (MENDEZ-FERRER et al., 2010; KUNISAKI et al., 2013; MORRISON e

SCADDEN, 2014). MSC is an essential component of HSC niche and displays an

important supportive role in hematopoiesis, especially on HSC and hematopoietic

progenitors (MENDEZ-FERRER et al., 2010).

Thus, in this study, we evaluated the alterations caused by PM on the regulatory

function of MSC on hematopoiesis. PM group received a hypoproteic diet and exhibited

anemia and leukopenia, as also reduction of blasts and hematopoietic precursors, as

described in previous studies (BORELLI et al., 2007; FOCK et al., 2007; FOCK,

BLATT, et al., 2010; FOCK, ROGERO, et al., 2010; SANTOS et al., 2017).

BM-MSC were isolated from control and malnourished mice and characterized

by CD44, CD49e and CD90.1 positive staining and absence of hematopoietic markers.

The immunophenotypic characterization and function of subtypes of MSC is still under

discuss. Since no MSC-specific marker has yet been identified, a set of at least 3

markers are used, the most frequently used ones being CD44, CD90, CD49e, CD29,

77

CD106, Sca-1, CD105, CD73 and CD271 (KOLF et al., 2007; KASSEM e ABDALLAH,

2008; MORIKAWA et al., 2009; KUCI et al., 2010).

Is has been reported that Nes+NG2+ and LepR+ MSC maintain HSC in a

quiescent stage by production of Ang-1 (KUNISAKI et al., 2013; ZHOU et al., 2015),

whereas Nes+ MSC and CAR cells (Sca-1-CD31-CD45-PDGFRa/b+ MSC) induce self-

renewal and proliferation on HSC and hematopoietic progenitors by SCF and CXCL-

12 release (NAGASAWA et al., 1996; SUGIYAMA e NAGASAWA, 2012).

Malnourished MSC expressed reduced levels of Igf1. IGF-1 plays an important

autocrine function on MSC by increasing proliferation rate and also inducing

osteoblastic and reducing adipogenic differentiation (YOUSSEF et al., 2017), which

corroborates the findings that PM increases adipogenic commitment of MSC (CUNHA

et al., 2013). In addition, decreased release of IFG-1 by malnourished MSC reduces

the expression of CXCR-4, the receptor of CXCL-12 (YOUSSEF et al., 2017). In spite

of malnourished MSC did not caused significative alterations in Cxcr4 expression on

c-Kit+ cells, the CXCL-12 synthesis was decreased. The activation of CXCR-4/CXCL-

12 axis mediates HSC quiescence and interrupts the progression of G1 to S cell cycle

phases, on the other hand, the lack of CXCR-4 impacts on increased levels of cyclin

D1 and enhances the G1 phase progression (CASHMAN et al., 2002; NIE et al., 2008).

In addition, PM decreased expression of Nes and Cspg4 (NG2) on MSC, and

also the production of Ang-1 and TGF-b and, consequently, PM can suppress cellular

self-renewal and prevent entry into the cell cycle (ARAI et al., 2004; WANG et al.,

2018). On the other hand, malnourished MSC boosted SCF synthesis. SCF is crucial

for hematopoiesis, since it mediates HSC survival and proliferation, as it directly

regulates the entry of hematopoietic cells into the cell cycle (LENNARTSSON e

RONNSTRAND, 2012). Altogether, these findings reveal that PM decreases the

capacity of MSC to induce HSC quiescence and, therefore, malnourished MSC are in

a pro-proliferative state.

Indeed, malnourished MSC exhibited capacity to enhance cell cycle progression

in control group, however PM group did not respond with the same magnitude. It has

been reported that PM impairs the entry of hematopoietic stem and progenitor cells

into cell cycle, by inducing the expression of the inhibitory proteins p21 and p27 and

by suppressing the induction proteins cyclin E, cyclin D1, Cdk2, Cdk4, and Cdc25a

(NAKAJIMA et al., 2014).

78

Additionally, the gene expression profile of c-Kit+ cells showed impairment of

differentiation-related genes and improvement of pluripotent genes in control group

after culture with supernatant from malnourished MSC, but malnourished group did not

respond to MSC stimuli as control group, which reinforces the idea of intrinsic

alterations in the c-Kit+ cells caused by PM. Malnourished MSC altered the expression

of the transcription factors Gata1 and Gata2, important transcriptions factors for

erythroid differentiation (MORIGUCHI e YAMAMOTO, 2014), and Gata3, the

transcription factor that regulates T cell lymphopoiesis (WAN, 2014).

To further confirm that malnourished MSC impairs the hematopoietic

differentiation into lineage-specific cells via soluble secreted factors, we quantified the

cell populations after cultures of hematopoietic cells and MSC supernatant from both

groups. Malnourished MSC boosted HSC, MPP and CMP populations in control group,

whereas reduced CLP, GMP and MEP, due to increased SCF concentrations. We

observed a decreased MEP population in PM group when control and malnourished

supernatants were compared. In addition to paracrine signaling of MSC, the contact

between HSC and MSC can regulate hematopoietic proliferation mediated via Notch

signaling pathway activation (GOTTSCHLING et al., 2007), however no differences

caused by malnourished MSC were observed in the quantification of hematopoietic

progenitors or in mature granulocytes and monocytes/macrophages.

Concluding, PM shifts MSC to a pro proliferative stage, alters the regulatory

function of MSC and promotes proliferation, although malnourished hematopoietic

cells cannot respond adequately to the stimuli from MSC. In addition, PM implicates in

loss of induction of lineage-specific differentiation, which leads to reduced

megakaryocytic-erythroid differentiation in malnourished mice

ACKNOWLEDGMENTS

We acknowledge the assistance of the School of Pharmaceutical Science Flow

Cytometry and Animal Care Cores.

This work was supported by Fundação de Amparo à Pesquisa do Estado de São Paulo

(FAPESP) grant (14/06872-2). R. A. Fock and P. Borelli are fellows of the Conselho

Nacional de Pesquisa e Tecnologia (CNPq).

The authors declare no competing financial interests.

79

Fig. 1. Results of nutritional parameters. Values for (A) diet consumption, (B) protein

consumption, (C) body weight variation, (D) total serum protein and (E) serum albumin

are expressed as mean ± SEM. Significant differences are illustrated by *(p≤0.05),

**(p≤0.01), ***(p≤0.001), ****(p≤0.0001). n represents the number of mice used in the

experiments.


1

2

3

4

5

Tota

l pro

tein

(g/d

L)

*

(C)

0 1 2 3 4 520

25

30

35

Weeks

Weig

ht va

riatio

n (g

) Control

Malnourished


1

2

3

4

5

Die

t con

sum

ptio

n(g/day/mouse)


0.2

0.4

0.6

Pro

tein

con

sum

ptio

n(g/day/mouse)

****

(D)

(A) (B)


0.5

1.0

1.5

2.0

2.5A

lbum

in (g

/dL

) **

(E)

0 1 2 3 4 520

25

30

35

Weeks

Wei

ght v

aria

tion

(g) Control

Malnourished

80

Table 1. Hematological evaluation Values for peripheral blood and bone marrow parameters are expressed as mean ±

SEM. Significant differences are illustrated by *(p≤0.05), **(p≤0.01), ***(p≤0.001),

****(p≤0.0001). n represents the number of mice used in the experiments.

Variables Control Group Malnourished Group

Peripheral blood parameters (n=20) (n=20)

Erythrocytes (106/mm3) 8,65 ± 0,22 7,90 ± 0,19 *

Hemoglobin (g/dL) 12,49 ± 0,27 11,05 ± 0,24 ***

Total leukocyte (/mm3) 2014 ± 89,98 841,2 ± 60,17 ****

Neutrophils (/mm3) 212,8 ± 20,03 113,8 ± 10,34 ***

Lymphocytes (/mm3) 1707,00 ± 21,80 653,10 ± 11,63 ****

Monocytes (/mm3) 36,55 ± 5,01 8,791 ± 2,54 ****

Platelets (x103/mm3) 536,1 ± 40,70 530,5 ± 39,90

Myelogram (n=5) (n=5)

Bone marrow cellularity (107/mm3) 3.01 ± 0,22 2,42 ± 0,13 ***

Blast cells (105/mm3) 4,3 ± 0,8 2,2 ± 0,1 ***

Granulocyte precursors (105/mm3) 10,6 ± 0,9 6,0 ± 0,9 ****

Band granulocytes (105/mm3) 19,0 ± 3,6 8,2 ± 1,1 ***

Polymorphonucleated granulocytes

(105/mm3)

134,2 ± 4,9 108,7 ± 2,3 ****

Monocytes (105/mm3) 2,2 ± 1,4 1,1 ± 0,6

Lymphocytes (105/mm3) 45,7 ± 10,1 23,9 ± 2,0**

Erythroblasts (105/mm3) 84,8 ± 6,9 56,5 ± 3,7 ****

81

Fig. 2. Characterization of mesenchymal stem cells from control and malnourished mice. (A) Heatmap of immunophenotypic characterization of MSC by

flow cytometry, results of CD34, CD45, CD11b, CD44, CD49e and CD90.1 are

expressed as log10 of positive cells (n=3). (B) Gene expression profile for mesenchymal

characterization in Control and Malnourished groups, results of gene expression of

Igf1, Icam1, Pdgf1, Eng, Mcam, Prom1, Nes, Cspg4 and Lepr are relative to 18s

expression and are expressed as mean and minimum to maximum values (n³8).

Significant differences are illustrated by *(p≤0.05). n represents the number of mice

used in the experiments.

(A) (B)

Control

Malnourished

Hematopoietic

markers

Mesenchymal

markers

CD34

CD45

CD11b

CD44

CD49e

CD90.1

-1

0

1

2

0 1 2 3

Lepr

Cspg4

Nes

Prom1

Mcam

Eng

Pdgf1

Icam1

Igf1

mR

NA

(rel

ativ

e to

18S

) Control Malnourished

*

**

82

Fig. 3. Regulatory status of hematopoiesis on MSC in PM. Results of MSC

production of (A) SCF, (B) TGF-b, (C) Ang-1, (D) CXCL-12 and (E) G-CSF by control

and malnourished groups are expressed as mean ± SEM (n=6). Results are expressed

as mean ± SEM. (F) Heatmap of gene expression of Angpt1, Cxcl12, Kitl, Il11, Il3,

Csf1, Csf2, Csf3 and Tgfb1 by control and malnourished groups, results are relative to

18s expression and are expressed as mean ± SEM (n³8). Significant differences are

illustrated by *(p≤0.05), **(p≤0.01). n represents the number of mice used in the

experiments.

Control Malnourished

Angpt1

Cxcl12

Kitl

Il11

Il3

Csf1

Csf2

Csf3

Tgfb1

0 1 2

**

***

(A)

(D) (E)

(C)(B) (F)


50

100

150

pg/mL

G-CSF

*


500

1000

1500

pg/mL

Ang-1

*


50

100

150

200

250

pg/mL

CXCL-12

*


20

40

60

pg/mL

TGF-β1

*


10

20

30

40

pg/mL

SCF

**

83

Fig. 4. Evaluation of the role of PM on MSC regulatory over gene expression in hematopoietic progenitor cells. Gene expression profile of c-Kit+ cells after

conditioned media with control MSC supernatant and conditioned media with

malnourished MSC supernatant of (A) pluripotency genes (Sox2, Nanog, and Pou5f1),

(B) chemokine receptors genes (Il3ra and Cxcr4), (B) lymphoid differentiation genes

(Ikzf3 and Gata3) and (D) myeloid differentiation genes (Gata1, Gata2, Nfe2, Spi1 and

Cebpa). n=3, results are relative to Gapdh expression and expressed as mean ± SEM.

Significant differences are illustrated by *(p≤0.05), **(p≤0.01), ***(p≤0.001). n

represents the number of mice used in the experiments.

(D)

(C)(B)(A)

Ikzf3 Gata30

1

2

3

4

mR

NA

(rel

ativ

e to

Gap

dh)

Expression of lymphoid differentiation genes***

***

***

*

Gata1 Gata2 Spfi1 Cebpa Nfe20.0

0.5

1.0

1.5

mR

NA

(rel

ativ

e to

Gap

dh)

Expression of myeloid differentiation genes

**

*

*

***

*** **

*

Sox2 Pou5f1 Nanog0

2

4

10

15m

RN

A (r

elat

ive

to Gapdh

)

Expression of pluripotency genes

Control + Cond. Control MSC Control + Cond. Malnourished MSC

Malnourished + Cond. Control MSC Malnourished + Cond. Malnourished MSC

***

***

*****

*** **

*

Il3ra Cxcr40

1

2

3

4

mR

NA

(rel

ativ

e to

Gap

dh)

Expression of cytokine receptors

CMP GMP MEP0.0

0.5

1.0

1.5

369

% o

f via

ble

cells

**

**

Control + Cond. Control MSC

Control + Cond. Malnourished MSC

Malnourished + Cond. Control MSC

Malnourished + Cond. Malnourished MSC

84

Fig. 5. Effects of conditioned media with control and malnourished EC supernatant over viability, cell cycle and differentiation. (A) BM-MNC viability and

apoptosis status after culture with culture media conditioned with control or

malnourished MSC supernatant. (B) Percentage of BM-MNC G0/G1 and S/G2/M cell

cycle phases after culture media conditioned with control or malnourished MSC

supernatant, n=3 each group. (C) Quantification of HSC and MPP populations after

culture of MNC and conditioned media with control MSC supernatant or conditioned

media with malnourished MSC supernatant. (D) Quantification of CLP, CMP, GMP and

MEP populations after culture of MNC and conditioned media with control MSC

supernatant or conditioned media with malnourished MSC supernatant. Gating

strategy is described in Supplemental Information (Fig. S1). Results are expressed as

mean ± SEM, n=3, each group. Significant differences are illustrated by *(p≤0.05),

**(p≤0.01), ***(p≤0.001) and ****(p≤0.0001). n represents the number of mice used in

the experiments.

(A) (B)

(C) (D)

CLP CMP GMP MEP0

1

2

3

% o

f via

ble

cells

********

*****

********

*****

********

HSC MPP0.00

0.05

0.10

0.15

0.20

0.25

% o

f via

ble c

ells

*** *

*** *

Viable cells Apoptotic cells0

20

40

60

80

100%

of c

ells

G0/G1 S/G2/M0

20

40

60

80

100

% of

cells





*

*****

******

CMP GMP MEP0.0

0.5

1.0

1.5

369

% o

f via

ble

cells

**

**





85

Fig. 6. Evaluation of the role of PM on MSC – c-Kit+ cells contact on lineage-specific differentiation in PM. (A) Evaluation of cell viability after c-Kit+ cells and MSC

from control and malnourished groups co-cultures. (B) Values of the quantification of

granulocytes and monocytes/macrophages after c-Kit+ cells and MSC from control and

malnourished groups co-cultures. (C) Values of the quantification of c-Kit+ cells after c-

Kit+ cells and MSC from control and malnourished groups co-cultures. (D) Values of

the quantification of CMP, GMP and MEP after c-Kit+ cells and MSC from control and

malnourished groups co-cultures. Results are expressed as mean ± SEM, n=3, each

group. Significant differences are illustrated by **(p≤0.01). n represents the number of

mice used in the experiments.

(A) (B)

(C)

(D)

Granulocytes Monocytes / Macrophages

0.0

0.5

1.0

1.51020304050

% o

f via

ble

cells

CMP GMP MEP0.0

0.5

1.0

1.5

369

% o

f via

ble

cells

**

**





c-Kit+ cells0

5

10

15

20

25

% o

f via

ble

cells

Viability0

20

40

60

80

100

% o

f via

ble

cells

CMP GMP MEP0.0

0.5

1.0

1.5

369

% o

f via

ble

cells

**

**





86

SUPPLEMENTARY MATERIAL

Fig. S1 Flow cytometry gates strategy. Forward Scatter x Side Scatter, single cells and

viable cells gates (a). Hematopoietic stem cells (R2) and hematopoietic multipotent progenitors

(R3) gates strategy (b). Common lymphoid progenitors (R2) gates strategy (c). Common

myeloid progenitors (R2), granule-monocytic progenitors (R3), and megakaryocytic-erythroid

progenitors (R4) gates strategy (d). Granulocytes (R2) and macrophages (R3) gates strategy

(e).

87

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5 CAPÍTULO III

A DIMINUIÇÃO DO RECEPTOR DE G-CSF NAS CÉLULAS PROGENITORAS GRANULOCÍTICAS CAUSA NEUTROPENIA NA DESNUTRIÇÃO PROTEICA

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Title: Impairment of G-CSF receptor on granulocytic progenitor cells causes neutropenia in protein malnutrition

Authors: Araceli Aparecida Hastreiter 1, Edson Naoto Makiyama1, Primavera Borelli 1,

Ricardo Ambrósio Fock 1*


Sciences, University of São Paulo, São Paulo, Brazil.





[email protected]

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ABSTRACT Hematopoiesis is a dynamic and controlled process in which all mature blood cells are

formed in the bone marrow (BM) as a result of an orchestrated mechanism of stimulus.

It is well known that protein malnutrition (PM) states are able to affect hematopoiesis

leading to severe leucopenia and reduced number of granulocytes, which act as the

first line of defense, being important to the innate immune response. Therefore, this

study aimed to elucidate some of the mechanisms involved in the impairment of

granulopoiesis in PM. Malnourished animals presented leucopenia associated with

reduced number of granulocytes and reduced percentage of granulocytic progenitors;

however, no differences were observed in the regulatory granulopoietic cytokine G-

CSF. Additionaly, the malnourished group presented impaired response to in vivo G-

CSF stimulus compared to control animals. PM was implicated in decreased ability of

c-Kit+ cells to differentiate into myeloid progenitor cells and downregulated STAT3

signaling. Furthermore, malnourished group exhibited impairment of G-CSF receptor

on granule-monocytic progenitors and this reduced expression was not completely

reversible with G-CSF treatment. Overall, this study implies that PM promotes intrinsic

alterations to hematopoietic precursors, which result in hematological changes, mainly

neutropenia, observed in peripheral blood in PM states.

Keywords: Protein malnutrition; granulocyte-colony stimulating factor, granulocytes,

granule-monocytic progenitors, G-CSF receptor.

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1. INTRODUCTION

Protein malnutrition (PM) modifies physiological responses, inducing cell

damage and commonly increases susceptibility to infection (KEUSCH, 2003;

SCRIMSHAW, 2010). PM can affect all systems and organs, but primarily affects

tissues with a high rate of cell turnover, such as hematopoietic tissue (BORELLI et al.,

2004).

Hematopoiesis is a dynamic and controlled process in which all mature blood

cells are formed in the bone marrow (BM) from a hematopoietic stem cell, which has

the ability of self-renewal and differentiation into hematopoietic progenitors and

hierarchically gives rise to lineage-specific progenitors, such as granule-monocytic

progenitors (GMP), which are able to produce granulocytes and monocytes

(WEISSMAN e SHIZURU, 2008). Granulocytes are produced in the bone marrow from

compromised granulocytic progenitors and once mature are released into the blood

and tissues, being cells important to the innate immune response, able to act as the

first line of defense in host resistance and wound healing (DAY e LINK, 2012;

NAUSEEF e BORREGAARD, 2014).

Granulocyte production is influenced and stimulated by granulopoietic

cytokines, especially granulocyte-colony stimulating factor (G-CSF) which is able to

increase granulopoiesis (YOSHIKAWA et al., 1995). G-CSF is a well-known

hematopoietic growth factor that stimulates the proliferation and differentiation of

myeloid progenitors, and all the biological activity of G-CSF is mediated through

interaction with a specific cell surface receptor, the G-CSF receptor (G-CSFr)

(MCKINSTRY et al., 1997). Moreover, G-CSF – G-CSFr signaling stimulates members

of the STAT family, especially STAT3 and this pathway has important regulatory

activity in granulopoiesis (MCLEMORE et al., 2001; TOUW e VAN DE GEIJN, 2007;

ZHANG et al., 2010).

It is well known that in PM states, when not associated with other diseases, the

number of granulocytic cells, especially neutrophils, are reduced, which predisposes

to higher susceptibility to infection (KEUSCH, 2003; SCRIMSHAW, 2010). Although

hematopoietic changes caused by PM have been described for a long time, little detail

is known about the mechanisms that affect the production and expansion of

hematopoietic cells. Thus, this study aimed to elucidate the effects of PM on

granulocytic cells production and expansion and the role of G-CSF in this regulation

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control.

2. MATERIAL AND METHODS

2.1 Animals and diet

Male C57Bl/6 mice, 45–60 days old, were obtained from the Production and

Experimentation Laboratory of the School of Pharmaceutical Sciences of the University

of São Paulo. The mice underwent an adaptation period (10 to 15 days), in which all

animals received normoproteic diet and water ad libitum until stabilization of body

weight. After this period, the animals were divided into two groups, which received

either normoproteic diet (Control group) or hypoproteic diet (Malnourished group)

(FOCK et al., 2007; FOCK, ROGERO, et al., 2010).

Both diets were prepared in-house, following the recommendations of the

American Institute of Nutrition for adult mice (REEVES et al., 1993). The protein source

used was casein (>85% protein). Both diets contained 100 g kg–1 sucrose, 80 g kg–1

soybean oil, 10 g kg–1 fiber, 2.5 g kg–1 choline bitartrate, 1.5 g kg–1 L-methionine, 40 g

kg–1 mineral mixture and 10 g kg–1 vitamin mixture. The control diet contained 120 g

kg–1 casein and 636 g kg–1 cornstarch, while the malnourishment diet contained 20 g

kg–1 casein and 736 g kg–1 cornstarch. With the exception of the protein and corn

starch content, the two diets were identical and isocaloric, providing 1716.3 kJ/100 g.

The final protein content of both diets was confirmed by the standard micro-Kjeldahl

method.

The period for the malnutrition induction was 35 to 40 days, and the weight and

feed intake of each animal were evaluated every 48 h (FOCK et al., 2007; FOCK,

ROGERO, et al., 2010). Protein consumption was calculated by the protein

concentration of the respective feed and the daily feed intake per animal. This project

was approved by the Animal Experimentation Ethics Committee of the School of

Pharmaceutical Sciences of the University of São Paulo.

2.2 In vivo G-CSF stimulus

Murine G-CSF (recombinant granulocyte-colony stimulating factor; Sigma

Chemical Company, St. Louis, MO, USA) was administered in the last four days of the

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period of malnutrition induction, intravenously via caudal vein in both Control and

Malnourished groups. The concentration administered was 8 µg/kg/day, previously

standardized (MOLINEUX et al., 1990; LORD et al., 1991; VINOLO et al., 2008). Sham

groups were performed with injection of sterile physiological saline solution. After the

6-hour period of the last administration, animals were euthanized and the samples

collected.

2.3 Blood

After the establishment of the experimental protocol, animals were anesthetized

and euthanized. Blood samples were collected for hematological evaluation and

plasma was separated by centrifugation (1000×g for 10 min at 4 oC). The

concentrations of plasmatic proteins, albumin and pre-albumin were determined by

standard methods (GORNALL et al., 1949; DOUMAS et al., 1971; HARRIS e KOHN,

1974). Cell blood counts were obtained by loading blood samples into ABX Micros

ABC Vet® equipment (Horiba ABX, Montpellier, France). The morphological and

leucocyte differential analyses were performed on blood smears stained by May-

Grünwald-Giemsa (Sigma Aldrich).

2.4 Bone marrow histology

Animals from the control and malnourished groups, stimulated or not with G-CSF,

had the sternum removed, which was immediately immersed in a 4% paraformaldehyde

fixative at room temperature for 24 h. The sternums were decalcified in 5% EDTA (pH

7.2) for one week. After decalcification, the sternums were processed by standard

histological techniques (paraffin-embedding). Five-micrometer sections of sternums

were stained by hematoxylin-eosin (H/E) and evaluated by conventional optical

microscopy.

2.5 Bone marrow cellularity and granulocytic lineage quantification by flow cytometry

Total BM cells obtained after femoral flushing with 10.0 mL of McCoy 5A (Sigma

Aldrich) supplemented with 10% fetal calf serum (Cultilab, Campinas, Brazil), 1%

penicillin and streptomycin (Sigma Aldrich), as described above, were used for

assessment of BM cellularity. BM cellularity was determined by counting obtained cells

using a Neubauer hemocytometer.

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To evaluate whether PM impairs the mature or progenitor granulocytic cells,

2 x 106 total BM cells were labeled with antibody cocktails and the cellular population

of granule-monocytic progenitors (GMPs, Lin-Il7r-c-Kit+Sca-1-CD34+CD16/32high) and

granulocytic cells (CD11b+Gr-1+) were quantified. Data were acquired on a FACS

Canto II (Becton Dickinson) and FlowJo® 10 software (TreeStar) was used for data

analysis. The antibodies used were CD3-PE (145-2C11), CD11b-PE (M1/70), CD11b-

PECy7 (M1/70), Ter119-PE (TER119), Ly6G-PE (RB6-8C5), CD19-PE (MB19-1),

Ly6A/E-PE (D7), CD127-PE (SB/199), CD34-FICT (RAM34), CD16/32-PECy7 (2.4G2)

AND c-Kit-APC (2B8), purchased from BD Biosciences. To establish negative controls,

we performed unstained and stained cells with fluorescence-minus-one (FMO) control

stain sets.

2.6 G-CSF quantification in bone marrow

After euthanasia, the femurs of each animal were removed. With needle and

syringe, BM cavities were flushed with 1.0 mL of McCoy 5A (Sigma Aldrich)

supplemented with 10% fetal calf serum (Cultilab, Campinas, Brazil), 1% penicillin and

streptomycin (Sigma Aldrich). BM flush was immediately centrifuged (350×g for 10 min

at 4 oC) and the supernatant was used for G-CSF quantification, determined by

Enzyme-Linked Immuno Sorbent Assay (ELISA) using commercially available kits

(Quantikine ELISA®, R&D Systems).

2.7 Ex vivo cell proliferation assay (CFU growth)

For ex vivo cell proliferation assay, BM c-Kit+ progenitor cells were isolated.

First, total BM cells were collected as described in section 2.6 from both femurs and

tibias, then mononuclear BM cells were separated by density gradient by Ficoll-

Histopaque technique (Sigma Aldrich). After that, the mononuclear cells were labeled

with anti-CD117 microbeads (Miltenyi Biotech Inc., Auburn, EUA) and c-Kit+ cells were

isolated using magnetic-activated cell sorting (MACS) following the manufacturer`s

instructions.

c-Kit+ cells (1 x 103 per well) were seeded and cultured in methylcellulose

semisolid medium (MethoCult M3630, StemCell Tech), supplemented with growth

factors (IL3, 0.1 ng/mL; EPO, 1UI; GM-CSF, 0.2 ng/mL; G-CSF, 2 ng/mL; and SCF,

50 ng/mL, Sigma Chemical Company, USA) Cells were incubated in a humidified

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atmosphere of 95% air, 5% CO2 at 37 °C for 14 days. After this period, the number of

clusters and colonies (PEREIRA et al., 2007) of CFU-GEMM (colony forming unit of

myeloid cells; granulocyte, erythrocyte, monocyte/macrophage, megakaryocyte) and

CFU-GM (colony forming unit of granule-monocytic cells) were counted using an

inverted microscope.

2.8 STAT3 expression by bone marrow c-Kit+ cells after G-CSF stimulus in vitro

The expression of STAT3 was evaluated in BM c-Kit+ cells by Western Blot

technique. BM c-Kit+ cells, obtained as described above, were seeded into 6-well

plates at a density of 5 × 105 cells per mL of culture medium and were stimulated or

not with 2 ng/mL murine G-CSF (Sigma Chemical Company, St. Louis, MO, USA) for

1 h. Subsequently, c-Kit+ cells were lysed with RIPA® buffer (Pierce, Rockford, USA)

containing protease and phosphatase inhibitors (0.5 mM PMSF, 50 mM NaF, 10 μg/mL

leupeptin, and 10 μg/mL aprotinin (Sigma Aldrich, St. Louis, USA). Protein

quantification was performed based on the Bradford method and a commercial kit

(BCATM protein assay kit®, Pierce, Rockford, USA) was used for this aim.

Subsequently, a sodium dodecyl sulfate polyacrylamide gel electrophoresis (10%) was

performed using 20 μg of protein sample followed by a polyvinylidene fluoride

membrane (PVDF®, Amersham Biosciences, Pittsburg, USA) transfer. A molecular

weight standard (BioRad, Philadelphia, USA) was used to compare separated

molecular weight fractions. Primary antibodies from Santa Cruz Biotechnology anti-

STAT3 (C-20, cat no. sc-482) and -pSTAT3 (Ser 727, cat no sc-8001-R) were diluted

in TBS-Tween buffer, respectively to 4:1,000 and 4:1,000, and incubated overnight.

Finally, membranes were incubated for 1 h with anti-IgG rabbit biotin- conjugated

secondary antibody (R&D Systems, Abingdon, UK) diluted to 1:10,000 in TBS-Tween

buffer. Immunoreactive bands were visualized using the ECL detection system®

(Amersham Biosciences, Pittsburg, USA) and images were captured using

ImageQuantTM 400® version 1.0.0 (Amersham Biosciences, Pittsburg, USA). For

standardization and quantification, images were analyzed using ImageQuant TL®

program (Amersham Biosciences, Pittsburg, PA, USA). Results were normalized to

the intensity of β-actin (Sigma-Aldrich, St. Louis, USA), which was diluted at 3:10,000

in TBS-Tween buffer.

2.9 Gene expression of G-CSFr on bone marrow c-Kit+ cells

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BM c-Kit+ cells were obtained as described above. Total RNA was obtained

using RNeasy extraction kit (Qiagen, USA) according to the manufacturer’s protocol.

Total RNA (50 ng) was reverse transcribed into cDNA using the High Capacity cDNA

reverse transcription kit (Applied Biosystems, USA). cDNA samples were then

amplified in the TaqMan® Fast Advanced master mix (Applied Biosystems) with

optimized concentrations of the primer set for Csf3r (Mm00432735_m1, Applied

Biosystems). The internal control used was Gapdh (Mm99999915_g1, Applied

Biosystems). The expression of Csf3r was evaluated by real-time PCR using

StepOnePlusTM (Applied Biosystems) and the relative gene expression quantification

was conducted according to the ΔΔCt method (LIVAK e SCHMITTGEN, 2001).

2.10 G-CSFr quantification on bone marrow granule-monocytic progenitor cells

To access the effect of PM on G-CSF receptor (G-CSFr) expression, which is

expressed at all maturation stages of purified myeloid cells, but in progenitor cells of

granulocytes have an important role in neutrophil production, 2 x106 total bone marrow

cells from both groups, injected or not with G-CSF, were labeled with antibody cocktails

as reported above and G-CSFr (CD114-AF488, #723806, R&D Systems) was

quantified on GMPs (Lin-Il7r-c-Kit+Sca-1-CD16/32high). Data were acquired on a FACS

Canto II (Becton Dickinson) and FlowJo® 10 software (TreeStar) was used for data

analysis.

2.11 Statistical analysis

All statistical analyses were performed with GraphPad Prism® 7 software

(GraphPad Software Inc., La Jolla, USA), and the data are expressed as mean ±

standard error of mean (SEM). Data sets passed through normality tests and were

analyzed by Student’s t-test. For data analysis of multiple comparisons among groups,

analysis of variance (2way ANOVA) plus Tukey's post hoc test. The level of significance

adopted was 95% (p < 0.05). Asterisks indicate a significant difference between

groups: *p < 0.05, **p < 0.01 and ***p < 0.001.

102

3. RESULTS 3.1 Nutritional status

Animals from Malnourished and Control groups exhibited similar food intake

during the period of malnutrition induction; however, malnourished animals had lower

protein intake due to the hypoproteic diet. As a consequence, malnourished animals

presented body weight loss and decreases in serum protein, albumin and pre-albumin

concentrations, as well as reduced erythrocyte count, hemoglobin concentration and

hematocrit (Table 1), characteristic of PM.

3.2 Malnutrition affects granulocytic cell count that G-CSF is not able to reverse

After the period of malnutrition, animals that received hypoproteic diet presented

peripheral leucopenia associated with reduction in granulocytic cells (Fig. 1 A and B)

and BM hypoplasia with reduced numbers of granulocytic cells (Fig. 1 C–E). As

observed in Fig. 2, malnourished animals presented with shrinkage of the marrow

hematopoietic space (Fig. 2 I–L) leading to hypocellular BM, and malnourished

animals stimulated with G-CSF did not increase significantly the number bone marrow

cells (Fig. 2 M–P). Animals from control group that received G-CSF showed increased

values of granulocytic cells both in BM compartment (Figs. 1 E and 2 H) as well as

peripherally (Fig. 1 B). However, malnourished animals that received G-CSF did not

respond to the stimulus with the same intensity when compared to control animals

(Figs. 1 B and 2 M–P).

3.3 Malnutrition affects GMP population and the response to G-CSF stimulus

Since PM caused medullary hypoplasia, we investigated if, specifically, the

GMP population was reduced. Malnourished animals showed reduced number of GMP

as well as granulocytic GR-1 positive cells in the BM, evaluated by flow cytometry.

Animals injected with G-CSF did not increase the percentage of GMP, as observed in

control animals, and although an increase in the percentage of GR-1 positive cells was

observed this was inferior in comparison to control group (Fig. 3).

3.4 Bone marrow G-CSF quantification

103

BM G-CSF was quantified in order to elucidate whether the alterations in BM

granulocyte and GMP populations are due to reduced G-CSF production. The G-CSF

was quantified in the BM of control and malnourished animals, injected or not with G-

CSF, and no differences in the G-CSF concentration between groups were observed

(Fig. 4A).

3.5 PM impairs colony-forming ability of c-Kit+ cells

To explain whether the response by malnourished animals to G-CSF stimulus

is inferior due to the lower number of GMP observed, the colony-forming assay was

performed. The colony-forming ability of c-Kit+ cells was determined using a

methylcellulose culture system containing SCF, EPO, GM-CSF, G-CSF and IL-3. The

results showed that the ability of c-Kit+ cells from malnourished animals to form CFU-

GM and CFU-GEMM was impaired in comparison to cells from control animals (Fig. 4B).

3.6 PM downregulates STAT3 signaling in c-Kit+ cells

Since G-CSF activates STAT3 signaling pathway on myeloid progenitors, the

expression of this transcription factor was assessed by Western Blot in BM c-Kit+ cells

with and without G-CSF stimulus in vitro. The results showed that the ratio between

phosphorylated (p-STAT3) and total STAT3 was significantly lower in malnourished

group when c-Kit+ cells were stimulated with G-CSF (Fig. 4C).

3.7 G-CSF receptor expression is impaired in protein malnutrition

Given that protein malnutrition compromises immune response and induces a

leucopenia hyporesponsive to G-CSF treatment, we investigated whether PM changes

G-CSF receptor (G-CSFr) expression. For that, the gene expression of G-CSFr (Csf3r)

was evaluated on BM c-Kit+ cells and the results showed that malnourished animals

exhibited reduced expression of Csf3r (Fig. 5A). Additionally, CD114 was evaluated

on GMP population by flow cytometry and the results showed reduced CD114 (G-

CSFr) expression in cells from malnourished animals (Fig. 5B). Malnourished animals

stimulated with G-CSF also showed reduced expression of G-CSFr in GMP population

(Fig. 5C-E). G-CSF treatment did not change the expression of G-CSFr in GMP of both

groups and G-CSFr expression remained lower in the malnourished GMP.

104

4. DISCUSSION

In this study, malnourished animals presented a quantitative reduction in

hematopoietic cells, especially the granulocytic lineage, in both BM and peripheral

blood compartments. This leucopenia can compromise both innate and acquired

immunity, as described in previous studies (FOCK et al., 2007; FOCK, BLATT, et al.,

2010; FOCK, ROGERO, et al., 2010; NAKAJIMA et al., 2014). Therefore, we

investigated some quantitative and qualitative alterations in hematopoietic progenitors

caused by PM, especially in GMP. The expansion of GMP is an important step for

emergency granulopoiesis response by promoting the production and maintenance of

granulocytic circulating cells pool, which act in the first defense line against infections

(DAY e LINK, 2012; NAUSEEF e BORREGAARD, 2014).

A previous report showed decreased hematopoietic stem cell (Lin-Sca-1+c-Kit+

- LSK) and progenitor cell (CD45+CD34+) populations in malnourished animals

(NAKAJIMA et al., 2014). However, here we first describe a specific reduction in GMP

(Lin-Il7r-c-Kit+Sca-1-CD34+CD16/32high) caused by PM, which explains, in part, why

there is a reduction of mature granulocytes, in the BM and peripheral blood.

This lack of progenitors is a consequence of the downregulation of some

mechanisms that drive hematopoietic stem cell and progenitor differentiation in a

lineage-specific manner. In this way, some cytokines and growth factors are

determinant for adequate hematopoiesis; G-CSF is essential for adequate

hematopoiesis, specifically granulopoiesis, since it is a potent regulator of the

development and function of granulocytes in vivo (LIESCHKE et al., 1994; LIU et al.,

1996; SEMERAD et al., 1999).

Additionally, G-CSF levels increase significantly during infections, promoting the

proliferation and differentiation of GMP and mobilization of mature granulocytes (LORD

et al., 1991). Knowing that, and once malnourished animals showed reduced

production of granulocytes, we first hypothesized that this reduction was due to a

reduced production of G-CSF in the hematopoietic production compartment; in other

words, reduced concentration of G-CSF in the bone marrow. However, when G-CSF

content in the bone marrow was measured, no differences were observed between

control and malnourished animals.

Moreover, we decided to stimulate animals with G-CSF. Clinically, G-CSF is

used to treat leucopenia due to suppression of bone marrow and also has the ability

105

to mobilize hematopoietic progenitor cells into the peripheral blood (LEVESQUE e

WINKLER, 2008; COOPER et al., 2011). Vinolo et al (2008) reported that

malnourished animals stimulated with G-CSF did not increase medullar or blood

granulocytes as well-nourished animals (VINOLO et al., 2008). In this study, we also

observed a discrete increase in mature granulocytes in malnourished animals after G-

CSF stimulus, but this increase was much lower than that observed in control group.

Moreover, the GMP percentage did not increase in malnourished group after the G-

CSF stimulus, presenting a completely opposite effect when compared to control

group, where an increased percentage of GMP was observed.

As we observed this decrease in myeloid progenitors and mature cells in the

BM caused by PM, we investigated the functionality of these cells in vitro to elucidate

whether there is less production of mature cells because there are fewer myeloid

progenitor cells. The ability of c-Kit+ cells from malnourished animals to produce CFU-

GEMM and CFU-GM was also impaired, indicating that there is also intrinsic

impairment in progenitor cells.

The growth-stimulatory effect of G-CSF-driven granulopoiesis depends on the

binding of G-CSF – G-CSFr and subsequent activation of STATs signaling pathways,

with a predominant activation of STAT3 (PANOPOULOS et al., 2006; TOUW e VAN

DE GEIJN, 2007; ZHANG et al., 2010). Therefore, we next investigated the STAT3

expression in BM c-Kit+ cells and our results showed decreased expression of

pSTAT3/STAT3 ratio after G-CSF stimulus in malnourished animals in comparison to

control animals. This inability of G-CSF to activate STAT3 in malnourished animals can

lead to an ineffective GMP proliferation and failure in the production of granulocytes

(TOUW e VAN DE GEIJN, 2007; MEHTA et al., 2015).

To try to understand, in part, this intrinsic impairment and the reason why the

malnourished animals did not have the same capacity to increase granulopoiesis after

G-CSF stimulus as did the control animals, we decided to evaluate the expression of

the G-CSF receptor (G-CSFr) in GMP, which are the myeloid progenitors that have the

main capacity to respond to G-CSF and consequently to induce granulopoiesis (CHEE

et al., 2013). In addition, the literature reports that animals that do not express G-CSFr

(G-CSFr-null) are severely neutropenic, with more pronounced reduction in peripheral

blood than in BM (LIESCHKE et al., 1994; LIU et al., 1996). In this way, interesting

results were observed, since the mRNA expression of G-CSFr (Csf3r) was reduced in

BM c-Kit+ cells from malnourished animals as well as the expression of G-CSFr on

106

GMP, which can explain, in part, why these animals presented a large reduction in

granulocytes in bone marrow and peripheral blood.

In addition, after stimulation with G-CSF we did not observe any changes in G-

CSFr expression in both groups in comparison to animals not stimulated, but again the

G-CSFr expression was reduced in malnourished animals. Extrapolating our results to

the clinic, patients suffering from PM (clinical and possibly subclinical) may not

successfully respond to G-CSF, due to the lower expression of G-CSFr.

In conclusion, this study evidenced that PM compromises GMP, affecting

granulopoiesis, and this effect is, in part, dependent on the reduced expression of G-

CSFr.

FUNDING

This work was supported by grants from the Fundação de Amparo a Pesquisa do

Estado de São Paulo – FAPESP (grant number: 2016/16463-8). Borelli P and Fock

RA are fellows of the Conselho Nacional de Pesquisa e Tecnologia (CNPq). Hastreiter

AA received scholarships from Coordenação de Aperfeiçoamento de Pessoal de Nível

Superior - Brasil (CAPES) and Conselho Nacional de Pesquisa e Tecnologia (CNPq).

107

Table 1. Nutritional evaluation. Values for nutritional parameters are expressed as

mean ± SEM. Significant differences between groups are illustrated by *(p<0,05) and

***(p<0,001). n represents the number of animals used in the experiments.

Variables Control Group Malnourished Group

Nutritional parameters (n=10) (n=10)

Food intake (g/day/animal) 3.43 ± 0.34 3.72 ± 0.09

Protein intake (g/day/animal) 0.41 ± 0.04 0.08 ± 0,002***

Body weight variation (%) 15.43 ± 1,46 -18.96 ± 0.81***

Total serum protein (g/dL) 4.76 ± 0.10 3.44 ± 0,12***

Serum albumin (g/dL) 1.95 ± 0.06 1.43 ± 0.07***

Serum pre-albumin (mg/dL) 9.34 ± 0.36 4.79 ± 0.31***

Erythrocytes (106/mm3) 9.08 ± 0.25 8.16 ± 0.19*

Hemoglobin (g/dL) 11.08 ± 0.19 9.82 ± 0.41*

Hematocrit (%) 37.2 ± 0.19 34.7 ± 0.89*

108

Figure 1. Values for granulocytic cell count. Results of peripheral leucocytes (A),

peripheral granulocytes (B), bone marrow total cells (C), bone marrow granulocyte

precursors (D) and bone marrow total granulocytes (E) are expressed as mean ± SEM

of control (C) animals, control animals injected with G-CSF (C + G-CSF), malnourished

(M) animals and malnourished animals injected with G-CSF (M + G-CSF). Significant

differences between groups are illustrated by *(p ≤ 0.05), **(p ≤ 0.01), ***(p ≤ 0.001);

n = 3, where n represents the number of animals used in the experiments.

109

Figure 2. Representative sections of bone marrow from control and malnourished

animals. Bone marrow biopsy section from a representative control animal, showing

normal cellularity with heterogeneous populations of cells at different stages of

maturation. Embedded in paraffin (HE stain, A x4; B x10; C x40; D x100). Bone marrow

biopsy section from a representative malnourished animal, showing severe

hypocellularity. Embedded in paraffin (HE stain, I x4; J x10; K x40; L x100). Bone

marrow biopsy section from a control animal stimulated with G-CSF, showing

increased number of cells specially granulocytes (HE stain, E x4; F x10; G x40; H x100;

arrows show granulocytes). Bone marrow biopsy section from a malnourished animal,

showing hypocellularity not complete reversible with G-CSF stimulus (HE stain, M x4;

N x10; O x40; P x100). Sections are representative of control and malnourished

animals stimulated or not with G-CSF.

110

Figure 3. Values for flow cytometry quantification of granule-monocytic precursors

(GMPs) (A), and mature granulocytic cells (GR-1+) (D). Results are expressed as mean

± SEM of control (C) animals, control animals injected with G-CSF (C + G-CSF),

malnourished (M) animals and malnourished animals injected with G-CSF (M + G-

CSF). Representative dot plots of GMP quantification in control (B) and malnourished

(C) animals. Representative dot plots of granulocyte cells GR-1+ quantification in

control (E) and malnourished (F) animals. Significant differences between groups are

illustrated by *(p ≤ 0.05), **(p ≤ 0.01), ***(p ≤ 0,001); n = 3, where n represents the

number of animals used in the experiments.

111

Figure 4. Results of bone marrow G-CSF quantification (A). Number of colony-forming

unit of myeloid cells (CFU-GEMM) and colony-forming units of granule-monocytic cells

(CFU-GM) of the clonogenic assays using bone marrow c-Kit+ cells of control (C) and

malnourished (M) animals (B). Western blot expression of p-STAT3 and STAT3 in c-

Kit+ cells stimulated or not with G-CSF. Results of p-STAT3 / STAT3 (C) are

represented in relation to the intensity of b-actin and expressed in arbitrary units. The

results are expressed as mean ± SEM (n = 3). n represents the number of animals

used in the experiments. Significant differences between groups are illustrated by **(p≤

0.01).

112

Fig. 5 Values of quantification of G-CSFr. Results of mRNA Csf3r (G-CSFr) in bone

marrow c-Kit+ cells from control and malnourished animals (A) are relative to Gapdh

expression and expressed as mean ± SEM (n = 6). Flow cytometry results of G-CSFr

in GMP population are expressed in mean of fluorescence intensity (B) and percentage

(C). Results are expressed as mean ± SEM of control (C) animals (n = 5), control

animals injected with G-CSF (C + G-CSF) (n = 3), malnourished (M) animals (n = 5)

and malnourished animals injected with G-CSF (M + G-CSF) (n = 3). Significant

differences between groups are illustrated by *(p ≤ 0.05) and ***(p ≤ 0.001); n

represents the number of animals used in the experiments. Representative dot plots

of G-CSFr quantification in control (D) and malnourished (E) animals in GMP

population with or without G-CSF stimulus.

113

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Elsevier Editorial System(tm) for Cytokine Manuscript Draft Manuscript Number: CYTO-18-705R1 Title: Impairment of G-CSF receptor on granulocytic progenitor cells causes neutropenia in protein malnutrition Article Type: Full length article Keywords: Protein malnutrition; granulocyte-colony stimulating factor, granulocytes, granule-monocytic progenitors; G-CSF receptor Corresponding Author: Professor Ricardo Fock, PhD Corresponding Author's Institution: University of Sao Paulo First Author: Araceli A Hastreiter Order of Authors: Araceli A Hastreiter; Edson N Makiyama; Primavera Borelli; Ricardo Fock, PhD Abstract: Hematopoiesis is a dynamic and controlled process in which all mature blood cells are formed in the bone marrow (BM) as a result of an orchestrated mechanism of stimulus. It is well known that protein malnutrition (PM) states are able to affect hematopoiesis leading to severe leucopenia and reduced number of granulocytes, which act as the first line of defense, being important to the innate immune response. Therefore, this study aimed to elucidate some of the mechanisms involved in the impairment of granulopoiesis in PM. Malnourished animals presented leucopenia associated with reduced number of granulocytes and reduced percentage of granulocytic progenitors; however, no differences were observed in the regulatory granulopoietic cytokine G-CSF. Additionaly, the malnourished group presented impaired response to in vivo G-CSF stimulus compared to control animals. PM was implicated in decreased ability of c-Kit+ cells to differentiate into myeloid progenitor cells and downregulated STAT3 signaling. Furthermore, malnourished group exhibited impairment of G-CSF receptor on granule-monocytic progenitors and this reduced expression was not completely reversible with G-CSF treatment. Overall, this study implies that PM promotes intrinsic alterations to hematopoietic precursors, which result in hematological changes, mainly neutropenia, observed in peripheral blood in PM states.

118

6 DISCUSSÃO FINAL

Os efeitos mais importantes da dieta sobre o organismo ocorrem a nível

molecular e podem ser tanto benéficos quanto prejudiciais, envolvendo diversos

órgãos em seus mais variados e complexos níveis de regulação (HIRSCH e EVANS,

2005). O menor consumo proteico pode desencadear uma série de fenômenos

fisiológicos para garantir a sobrevivência, mas que são deletérios a longo prazo

(COZZOLINO e COMINETTI, 2013).

Os constituintes da dieta, tanto macro quanto micronutrientes, participam da

regulação da expressão gênica em resposta a alterações nutricionais (CORTHESY-

THEULAZ et al., 2005). Por exemplo, a DP/DPE modifica, epigeneticamente,

mecanismos de controle do estresse, que resultam em alterações na síntese de

adrenalina, noradrenalina, cortisol e hormônio adrenocorticotrófico. Estas alterações

metabólicas podem causar malformação vascular e desencadear alterações

permanentes no metabolismo intermediário, levando à menor oxidação de gordura e

danos na síntese muscular e óssea e, portanto, a DP não causa consequências para

o organismo somente durante sua instalação (COZZOLINO e COMINETTI, 2013). Em

longo prazo, a DP pode favorecer o desenvolvimento de doenças crônicas não

transmissíveis, como diabetes, hipertensão e obesidade, sendo que já está bem

estabelecido na literatura a correlação entre DP/DPE na infância e obesidade na idade

adulta, principalmente em mulheres (FLORENCIO et al., 2008; FERREIRA et al.,

2009).

Neste trabalho, o modelo experimental utilizado induziu a desnutrição proteica

a partir de uma dieta hipoproteica. As dietas normoproteica e a hipoproteica utilizadas

são isocalóricas e com teores de vitaminas, ácidos graxos e sais minerais similares

para fornecer uma dieta adequada, restringindo-se apenas a oferta de caseína

(REEVES et al., 1993; REEVES, 1997). O período de indução da desnutrição utilizado

neste trabalho (5 semanas) equivale a dois anos de idade para o homem (QUINN,

2005), e, dessa forma, pode ser considerado equivalente a dois anos de alterações

na dieta e, portanto, considerado como uma afecção crônica.

Como consequência da instalação da DP, os animais que receberam dieta

hipoproteica apresentaram redução de peso corpóreo, devido ao aumento do

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catabolismo muscular observado na privação proteica, em que o organismo tende a

utilizar aminoácidos provenientes da musculatura esquelética e, em menor proporção,

da musculatura lisa (HUANG e FRAKER, 2003; ALVES et al., 2008; MALAFAIA et al.,

2009).

Em virtude da menor ingestão proteica pelos animais do grupo desnutrido,

observamos, conforme esperado, redução nas concentrações séricas de proteínas

totais, albumina e pré-albumina. Assim, a perda de aproximadamente 20% do peso

corpóreo inicial, associados aos valores das concentrações de proteínas totais e

albumina séricas e ao hemograma, mostraram-se bons indicadores da instalação da

DP.

Observamos neste trabalho um quadro hematológico condizente com a

instalação da DP, decorrente de hipoplasia medular e que corrobora os demais

trabalhos do grupo, em que há diminuição da celularidade medular, redução do

número de hemácias, redução na porcentagem do volume do hematócrito e baixa

concentração de hemoglobina no grupo desnutrido. Adicionalmente, foi observado

redução quantitativa expressiva no número leucócitos tanto no compartimento

periférico quanto no central (BORELLI et al., 1995; BORELLI et al., 2004; BORELLI et

al., 2007; XAVIER et al., 2007; FOCK, ROGERO, et al., 2010; FOCK et al., 2012; DOS

SANTOS et al., 2017). O número de plaquetas foi semelhante nos dois grupos

avaliados e está de acordo com dados da literatura descritos desde 1978 (FRIED et

al., 1978).

Foi descrito anteriormente que a DP diminui a população de células tronco e

progenitoras hematopoéticas Lin-Sca-1+c-Kit+ (LSK) (NAKAJIMA et al., 2014),

entretanto esta população celular é heterogênea e abrange somente as CTH, MPP e

CLP. Demonstramos neste trabalho que a DP provoca também a redução dos

progenitores mieloides CMP, GMP e MEP, bem como evidenciamos redução,

isoladamente, das populações de CTH, MPP e CLP, que foi acompanhada da

supressão da expressão dos genes de fatores de transcrição que conduzem à

diferenciação da CTH em progenitores linhagem-específicos.

Os processos de proliferação e diferenciação da CTH e dos progenitores

hematopoéticos são rigorosamente controlados por uma série de fatores de

transcrição. Os principais fatores de transcrição relacionados à diferenciação linfoide

são GATA3 e IKZF3 e à diferenciação mieloide são GATA1, GATA2, NF-E2, PU.1 E

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C/EBPa. Estes fatores de transcrição não atuam apenas em um tipo específico de

célula progenitora hematopoiética, mas, de acordo com sua ação mais proeminente,

podemos inferir que IKZF3 (Ikaros family zinc finger 3) conduz à diferenciação de

linfócitos B, enquanto que GATA3 induz a diferenciação de linfócitos T (NAKAJIMA,

2011). GATA1 é um regulador essencial no desenvolvimento da linhagem eritroide,

por controlar a sobrevivência e diferenciação dos eritroblastos. GATA2 também está

envolvido na diferenciação eritroide, mas, mais importante, está relacionado à

capacidade de autorrenovação da CTH e dos MPP (IWASAKI et al., 2006). Já NF-E2

induz a diferenciação megacariocítica, enquanto PU.1 e C/EBPa controlam diferentes

estágios de diferenciação granulocítica (IWASAKI et al., 2006; MONTICELLI e

NATOLI, 2017).

A diminuição das CTH e dos progenitores hematopoéticos é consequência de

da parada do ciclo celular causada pela DP, observada neste trabalho e em

concordância com estudos anteriores que relataram maior porcentagem de células

LSK nas fases G0/G1 do ciclo celular (BORELLI et al., 2009; NAKAJIMA et al., 2014)

e que indicam que a DP pode aumentar o número de CTH em estado de quiescência.

Fisiologicamente, a maioria das CTH está em estado quiescente, ou seja, na

fase G0 do ciclo celular (ROSSI et al., 2007). Dessa forma, a regulação da progressão

da fase G0 para G1 do ciclo celular pelo complexo ciclina D –Cdk4/6 é um fator

determinante na regulação da quiescência (PASSEGUE et al., 2005).

A manutenção da quiescência é essencial para a manutenção do pool de CTH,

visto que em baixa atividade metabólica há diminuição do stress oxidativo e,

consequentemente, há menores níveis intracelulares de espécies reativas de

oxigênio e nitrogênio (ARAI e SUDA, 2007), que por sua vez, induzem a diferenciação

das CTH (NOGUEIRA-PEDRO et al., 2014).

Dessa forma, a perda da quiescência leva ao comprometimento da

autorrenovação e pode resultar no esgotamento das CTH (ORFORD e SCADDEN,

2008). Adicionalmente, observamos que a DP suprimiu a expressão dos genes de

pluripotência, que, igualmente, compromete a autorrenovação das CTH, além de

diminuir a capacidade de recuperar o tecido hematopoiético.

A produção de células sanguíneas em um padrão constante depende do

microambiente hematopoético. Funcionalmente, as CTH são controladas por uma

combinação de fatores intrínsecos e por mecanismos externos responsáveis por

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regular sua proliferação, diferenciação e, por fim, na quiescência celular (PIETRAS et

al., 2011). Adicionalmente, o microambiente regula a localização e fisiologia das

células hematopoéticas, bem como a taxa de liberação de células maduras da MO

para o sangue periférico (MAYANI et al., 1992; VITURI et al., 2000).

Dessa forma, averiguamos nesta pesquisa se a DP acarreta alterações no

microambiente medular que promovam as alterações hematológicas descritas.

Primeiramente, investigamos se o microambiente medular de animais desnutridos

oferece suporte adequado à proliferação celular in vivo. Para tanto, realizamos

transplantes de mielo-monoblastos leucêmicos em animais sadios e desnutridos, em

que observamos menor taxa de proliferação e maior percentual das células

transplantadas nas fases G0/G1 do ciclo celular nos animais desnutridos. Este fato

nos leva a inferir que há alterações no microambiente medular que interrompem a

progressão do ciclo celular, confirmando nossa hipótese de que a MO não suporta a

hematopoese numa situação de DP.

Atualmente, estão descritos como nichos hematopoéticos o nicho endosteal e

o nicho perivascular, cada um com suas características distintas. Entretanto, estes

nichos ainda são pouco compreendidos, pois pouco se entende sobre os

mecanismos que regulam a formação do microambiente medular responsável pelo

controle da hematopoese, principalmente se considerarmos a reduzida quantidade

de informações acerca de sua localização in situ. Além disso, a literatura mostra

dados conflitantes acerca da função exata de cada um deles in vivo e a determinação

na íntegra dos componentes dos nichos não é consensual entre os trabalhos.

Nos últimos anos, o foco dos estudos que visam compreender a regulação do

microambiente sobre a hematopoese, foi o microambiente perivascular. Uma vez que

as CTH se localizam a cerca de 2 a 5 células dos vasos medulares e

aproximadamente 10 células da região da metáfise óssea, as CTH podem residir tanto

no endósteo como longe deste, mas elas sempre serão influenciadas pela vasculatura

(ELLIS et al., 2011). Contudo, devido à heterogeneidade de células envolvidas na

regulação desse microambiente, o nicho perivascular mostra-se bastante complexo,

o que se traduz em uma grande dificuldade para estudar a regulação do nicho in vivo

e/ou reproduzi-lo in vitro.

As CTM são um componente essencial do microambiente hematopoético e

desempenham um importante papel de suporte à hematopoese, sobretudo à

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regulação das CTH e dos progenitores hematopoéticos (GARCIA-GARCIA et al.,

2015). Em determinadas condições fisiológicas ou fisiopatológicas, as CTM podem

tanto se diferenciar em células especializadas, como osteoblastos, adipócitos, células

endoteliais e reticulares, quanto manterem o seu estado indiferenciado, secretando

fatores de crescimento e citocinas (SHI, 2012).

Isolamos e cultivamos CTM medulares de animais dos grupos controle e

desnutrido pela metodologia clássica descrita por Friedenstein, que utiliza a

propriedade física de aderência ao plástico (FRIEDENSTEIN et al., 1976). A

caracterização destas células ainda é complexa, pois, além de não haver um

marcador específico para as CTM, a expressão dos marcadores pode variar conforme

a espécie e mesmo entre as diferentes linhagens de camundongos (ANJOS-AFONSO

e BONNET, 2011; BOXALL e JONES, 2012). Atualmente, as CTM são caracterizadas

por uma combinação de critérios físicos e fenotípicos, além de propriedades

funcionais, como a confirmação de sua multipotencialidade, utilizando testes de

diferenciação osteogênica, adipogênica e condrogênica. Devido à sua extensa

plasticidade, a existência de subpopulações de CTM justificaria a sua

multipotencialidade e variedade fenotípica (PHINNEY, 2007; BOXALL e JONES,

2012). Os resultados de caracterização das CTM obtidos por imunofenotipagem por

citometria de fluxo e pelos testes de diferenciação mostraram que o isolamento e a

expansão celulares realizados foram efetivos.

Observamos que a DP promove alteração funcional nas CTM, visto que os

animais desnutridos sintetizaram menor quantidade de CXCL-12 e maior quantidade

de SCF que camundongos bem nutridos (HASTREITER, 2014). A CXCL-12 (ou SDF-

1) é a principal quimiocina que promove a mobilização da CTH do nicho endosteal

para o nicho perivascular na MO, de forma que quanto menor a quantidade de CXCL-

12, maior a mobilização e menos quiescente está a CTH. Sua interação com seu

receptor CXCR-4 regula não somente a movimentação da CTH, mas também a

adesão e sobrevivência celulares através de modulação do ciclo celular (NERVI et al.,

2006; GREENBAUM et al., 2013). O SCF é uma molécula que ativa o receptor tirosina

quinase c-Kit. Esta ativação é crucial para a hematopoese, visto que media a

sobrevivência, migração e proliferação celulares (LENNARTSSON e RONNSTRAND,

2012).

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Para elucidar como as células hematopoéticas respondem aos estímulos pró-

proliferativos provenientes da CTM, avaliamos como as MNC se comportam frente a

esses estímulos, através de culturas celulares com sobrenadantes provenientes de

CTM de animais controles e desnutridos, para avaliar efeitos parácrinos, e através de

sistemas de co-cultura, para avaliar o efeito do contato célula-célula. Observamos que

as CTM regulam a proliferação celular, conforme visto nos resultados da avaliação do

ciclo celular. As MNC dos animais controle respondem aos estímulos das CTM

controle, porém o grupo desnutrido mostrou-se mais quiescente. Ao avaliar as fases

do ciclo celular no grupo desnutrido, notamos que não há diferença ao comparar as

MNC ex vivo e após os tratamentos com CTM dos grupos controle ou desnutrido,

indicando que a parada maturativa destas células não é devido aos efeitos parácrinos

das CTM.

Além das CTM, o nicho perivascular apresenta uma heterogeneidade de

células que podem modular a hematopoese, como as CE. A identificação e o papel

de tipos distintos de CE ainda não são completamente compreendidos, mas estudos

com CE de diferentes fenótipos evidenciam sua importância na modulação da

hematopoese (DING et al., 2012; SASINE et al., 2017; KENSWIL et al., 2018).

Estudos recentes sugerem que as CE arteriolares são as principais produtoras de

SCF e promovem a manutenção das CTH (XU et al., 2018), enquanto as CE

sinusoidais controlam a diferenciação hematopoiética e a liberação de células

maduras para o sangue periférico, mas a distinção entre essas células não está

totalmente estabelecida e poucos estudos in vivo foram realizados. Além disso, ainda

não se sabe se as CE modulam a hematopoese apenas através de sinais parácrinos

ou se o contato célula-célula é indispensável.

As CE medulares são definidas como células CD144+ CD31+(DING et al.,

2012), com ausência de marcadores de células hematopoiéticas jovens. No presente

trabalho, obtivemos CE CD144+ CD31+ através da indução da transdiferenciação de

MSC em CE e observamos que a DP não prejudicou este processo. Embora não

tenham sido observadas alterações fenotípicas, a função destas células está

alterada, como evidenciado na quantificação de Ang-1, CXCL-12, SCF, IL-11 e G-

CSF.

Talvez o efeito mais significativo da DP na modulação da CE sobre a

hematopoese esteja relacionado ao ciclo celular. As CE de camundongos

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desnutridos produziram menos SCF que o grupo controle e o SCF regula diretamente

a entrada de células hematopoiéticas no ciclo celular (LENNARTSSON e

RONNSTRAND, 2012). A deleção condicional de SCF em células endoteliais e

perivasculares LepR+, mas não em osteoblastos e em células mesenquimais

nestina+, leva ao esgotamento de CTH (DING et al., 2012). Como resultado dessa

diminuição de SCF, uma quantidade menor de CTH foi detectada em culturas

condicionadas pela CE dos camundongos desnutridos. Além disso, as CE do grupo

desnutrido propiciaram o aumento da expressão de Cxcr4 nas células c-Kit+ de

camundongos desnutridos. A ativação da via CXCR-4/CXCL-12 promove a

quiescência celular, por diminuir a síntese de ciclina D1 e, consequentemente,

interromper a progressão da fase G1 para S do ciclo celular (CASHMAN et al., 2002;

NIE et al., 2008)

Visto que a DP induziu parada do ciclo celular das células hematopoéticas e

dos mielo-monoblastos leucêmicos transplantados de forma similar, nós avaliamos o

impacto parácrino das CE na expressão de proteínas de indução e inibitórias do ciclo

celular in vitro nas células C1498 (mielo-monoblastos leucêmicos). A DP induz a

expressão das proteínas inibitórias p21 e p27 e, por outro lado, suprime as proteínas

de indução ciclina E, ciclina D1, Cdk2, Cdk4 e Cdc25a (NAKAJIMA et al., 2014).

Observamos que as CE de camundongos desnutridos diminuíram a expressão de

ciclina E e D1 (Ccne1 e Ccnd1, respectivamente) e aumentaram a expressão de p21

e p27 (Cdkn1a e Cdkn1b, respectivamente) nos mielo-monoblastos in vitro, indicando

que a indução de quiescência nas CTH observada na DP é, pelo menos em parte,

devido a um efeito inibitório do ciclo celular pelas CE.

Não encontramos evidências que reforcem a participação da CE na indução

da linfopenia observada na DP. Embora as CE de camundongos desnutridos tenham

aumentado a expressão de CXCR-4 in vitro nas células c-Kit+ e que essa expressão

seja relevante para a diferenciação de MPP em CLP, observamos raros CLP após as

culturas condicionadas com CE, bem como supressão da expressão dos genes que

controlam a diferenciação dos progenitores linfoides (Gata3 e Ikzf3).

Observamos que as CE aumentam a síntese de IL-11 na DP, que

indiretamente aumenta a megacariocitopoese e a eritropoese e, em menor extensão,

a linfopoese através de um efeito sinérgico com outras citocinas e fatores de

crescimento, como IL-3, IL-4 e SCF (WADHWA e THORPE, 2008).

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Demonstramos que a DP induz efeitos parácrinos pela CE, bem como de

contato célula-CE, que redirecionam a diferenciação granulocítica para

megacariocítica e eritroides, apesar da maior síntese de G-CSF in vitro pelas CE dos

camundongos desnutridos.

A granulopoese é extremamente dependente de G-CSF, e em menor

extensão, de fator estimulador de colônias mononócitos e granulócitos (GM-CSF). O

G-CSF controla não somente a proliferação de células granulocíticas, mas também

a função dos neutrófilos maduros pela regulação direta da expressão de Sfpi1 (PU.1),

que codifica a principal molécula de adesão – CD11b – nos neutrófilos (LIESCHKE

et al., 1994; LIU et al., 1996; SEMERAD et al., 1999). As CE de camundongos

desnutridos aumentaram a expressão de Spi1 nas células progenitoras

hematopoéticas in vitro, porém não observamos aumento na produção de

granulócitos.

Dessa forma, inferimos que a neutropenia observada na DP não ocorre por

intermédio das CE e investigamos outras possíveis causas do comprometimento

granulocíticos. Em seguida, investigamos se a menor produção de G-CSF pelas CE

poderia refletir em menor quantidade de G-CSF na medula ex vivo. As CE são as

maiores produtoras de G-CSF in vivo, porém outras células do nicho podem produzi-

lo. Não encontramos diferença na quantificação de G-CSF no lavado medular entre

camundongos controle e desnutrido, por este motivo suspeitamos de alterações

intrínsecas nos progenitores hematopoéticos que podem conduzir à neutropenia.

Para tanto, decidimos estimular animais com G-CSF. Clinicamente, o G-CSF

é utilizado para tratar leucopenia devido à supressão da medula óssea e tem a

capacidade de mobilizar células progenitoras hematopoéticas para o sangue

periférico (COOPER et al., 2011; CHEE et al., 2013), e embora um discreto aumento

de granulócitos maduros tenha sido observado no grupo desnutrido, esse aumento

foi muito menor do que o observado em camundongos controle.

No entanto, o percentual de GMP não aumentou em animais desnutridos após

o estímulo com G-CSF, apresentando um efeito completamente oposto quando

comparado aos animais controle, onde foi observado um aumento na porcentagem

de GMP. Como observamos esta diminuição em progenitores mieloides e células

maduras na medula óssea causada por PM, nós investigamos a funcionalidade

destas células in vitro para elucidar se há menos produção de células maduras

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porque há menos células progenitoras mieloides. A capacidade de c-Kit + células de

animais desnutridos para produzir CFU-Mix e CFU-GM também foi prejudicada,

indicando que há também um comprometimento intrínseco nas células progenitoras.

Para tentar entender, em parte, esse comprometimento intrínseco e a razão

pela qual os camundongos desnutridos não apresentaram a mesma capacidade de

aumentar a granulopoese após o estímulo com G-CSF, decidimos avaliar a

expressão do receptor G-CSF (G-CSFr) em GMP, que são os progenitores mieloides

que têm a capacidade principal de responder ao G-CSF e consequentemente induzir

a granulopoiese (CHEE et al., 2013). Além disso, a literatura relata que camundongos

que não expressam G-CSFr (G-CSFR-null) são gravemente neutropênicos, com

redução mais pronunciada no sangue periférico do que na BM (SEMERAD et al.,

1999; LEVESQUE e WINKLER, 2008). Resultados interessantes foram obtidos,

mostrando que há expressão reduzida de G-CSFr em GMP em camundongos

desnutridos, o que pode explicar, em parte, por que esses animais apresentaram uma

grande redução de granulócitos na medula óssea e no sangue periférico. Além disso,

após estimulação com G-CSF, não observamos quaisquer alterações na expressão

de G-CSFr em ambos os grupos em comparação com animais não estimulados, mas

novamente a expressão de G-CSFr foi reduzida em animais desnutridos.

Extrapolando os nossos resultados para a clínica, os pacientes que sofrem de DP

(clínica e possivelmente subclínica) podem não responder com sucesso ao G-CSF,

devido à menor expressão de G-CSFr.

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7 CONCLUSÕES

• A DP compromete a hematopoese, reduzindo as populações de CTH e dos

progenitores hematopoéticos (MPP, CLP, CMP, GMP e MEP), bem como suprime a

expressão gênica de fatores de transcrição de pluripotência (Sox-2, Nanog e Oct-4) e

de diferenciação (Gata1/2/3, NF-E2, PU.1, C/EBP-a e IKZF3);

• O microambiente medular de camundongos desnutridos não sustenta a

hematopoese in vivo;

• As CTM apresentam-se em estado pró-proliferativo in vitro, devido à maior

síntese de SCF e menor síntese de Ang-1 e TGF-b1, bem como modulam a

diferenciação megacariocítica-eritróide, na DP;

• As CE induzem parada no ciclo celular das células tronco e progenitoras

hematopoéticas in vitro;

• A DP compromete a granulopoese, em parte, devido à redução da expressão

de G-CSFr nos progenitores granulocíticos.

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ANEXOS

ANEXO I – Protocolo da Comissão de Ética no Uso de Animais

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ANEXO II – Ficha do Aluno

- Sistema Administrativo da Pós-Graduação

Universidade de São PauloFaculdade de Ciências Farmacêuticas

Documento sem validade oficialFICHA DO ALUNO

9136 - 7913950/2 - Araceli Aparecida Hastreiter

Email: [email protected] de Nascimento: 05/11/1980Cédula de Identidade: RG - 3.633.127 - SCLocal de Nascimento: Estado de Santa CatarinaNacionalidade: Brasileira

Graduação: Farmacêutico - Universidade Federal de Santa Catarina - Santa Catarina - Brasil -2003

Mestrado: Mestra em Ciências - Área: Análises Clínicas - Faculdade de CiênciasFarmacêuticas - Universidade de São Paulo - São Paulo - Brasil - 2014

Curso: DoutoradoPrograma: Farmácia (Fisiopatologia e Toxicologia)Área: Análises ClínicasData de Matrícula: 01/10/2014Início da Contagem de Prazo: 01/10/2014Data Limite para o Depósito: 29/01/2019

Orientador: Prof(a). Dr(a). Ricardo Ambrosio Fock - 01/10/2014 até o presente. Email:[email protected]

Proficiência em Línguas: Inglês, Aprovado em 01/10/2014

Prorrogação(ões): 120 diasPeríodo de 01/10/2018 até 29/01/2019

Data de Aprovação no Exame deQualificação: Aprovado em 23/11/2016

Estágio no Exterior: Miami University, Estados Unidos da América - Período de 09/04/2018 até09/05/2018

Data do Depósito do Trabalho:Título do Trabalho:

Data Máxima para Aprovação daBanca:Data de Aprovação da Banca:

Data Máxima para Defesa:Data da Defesa:Resultado da Defesa:

Histórico de Ocorrências: Primeira Matrícula em 01/10/2014Prorrogação em 15/08/2018

Aluno matriculado no Regimento da Pós-Graduação USP (Resolução nº 6542 em vigor de 20/04/2013 até 28/03/2018).Última ocorrência: Prorrogação em 15/08/2018Impresso em: 19/01/2019 23:36:46

148

- Sistema Administrativo da Pós-Graduação

Universidade de São PauloFaculdade de Ciências Farmacêuticas

Documento sem validade oficialFICHA DO ALUNO

9136 - 7913950/2 - Araceli Aparecida Hastreiter

Sigla Nome da Disciplina Início Término CargaHorária

Cred.Freq.Conc.Exc.Situação

FBC5792-3/2 Tópicos em Análises Clínicas III 03/03/2015 16/06/2015 15 1 87 A N Concluída

BIO5788-3/1

Inglês em Ciência (Instituto de Biociências -Universidade de São Paulo) 04/03/2015 16/06/2015 120 0 - - N

Pré-matrículaindeferida

VCI5790-1/1

Modelos Animais para Terapia CelularExperimental (Faculdade de MedicinaVeterinária e Zootecnia - Universidade deSão Paulo)

10/04/2015 22/05/2015 30 2 100 A N Concluída

FBA5728-3/11 Aprimoramento Didático 14/04/2015 11/05/2015 60 0 - - N

Pré-matrículaindeferida

FBC5734-3/1

Aplicações da Citometria de Fluxo emModelos Experimentais 03/08/2015 09/08/2015 30 2 100 A N Concluída

FBC5766-4/2 Tópicos em Análises Clínicas IV 04/08/2015 16/11/2015 15 1 90 A N Concluída

MPT5793-1/5

Citogenômica I (Faculdade de Medicina -Universidade de São Paulo) 02/09/2015 27/10/2015 120 8 100 A N Concluída

MPT5778-3/2

Patometria I (Faculdade de Medicina -Universidade de São Paulo) 08/09/2015 02/11/2015 120 8 83 B N Concluída

VCI5785-2/1

Tópicos em Cultura Celular, com Ênfase emCultura Primária de Células Tronco(Faculdade de Medicina Veterinária eZootecnia - Universidade de São Paulo)

23/11/2015 29/11/2015 30 2 100 A N Concluída

FBC5748-4/2

Trabalhos Científicos: da Elaboração àPublicação 05/04/2016 17/05/2016 60 4 75 A N Concluída

VNP5733-5/3

Preparação Pedagógica - Nutrição eProdução Animal (Faculdade de MedicinaVeterinária e Zootecnia - Universidade deSão Paulo)

12/05/2016 23/06/2016 60 0 - - N Matrículacancelada

FBA5728-4/3 Aprimoramento Pedagógico 16/08/2016 12/09/2016 60 4 100 A N Concluída

BMI5902-1/1

Interações Imuno-Metabólicas (Instituto deCiências Biomédicas - Universidade de SãoPaulo)

03/08/2017 09/11/2017 60 0 - - N Matrículacancelada

ICB5752-1/3

Como Comunicar Sua Ciência: Melhorandoa Oratória e a Empatia com o Público(Instituto de Ciências Biomédicas -Universidade de São Paulo)

16/10/2017 29/10/2017 30 2 100 A N Concluída

Créditos mínimos exigidos Créditos obtidosPara exame de qualificação Para depósito de tese

Disciplinas: 0 20 34

Estágios:Total: 0 20 34

Créditos Atribuídos à Tese: 167

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