Monocytes, macrophages, dendritic cells and neutrophils: an update on lifespan kinetics in health and disease

Summary

Phagocytes form a family of immune cells that play a crucial role in tissue maintenance and help orchestrate the immune response. This family of cells can be separated by their nuclear morphology into mononuclear and polymorphonuclear phagocytes. The generation of these cells in the bone marrow, to the blood and finally into tissues is a tightly regulated process. Ensuring the adequate production of these cells and their timely removal is key for both the initiation and resolution of inflammation. Insight into the kinetic profiles of innate myeloid cells during steady state and pathology will permit the rational development of therapies to boost the production of these cells in times of need or reduce them when detrimental.

Patel et al., update our understanding regarding human phagocyte kinetics in health and disease.

Abbreviations

CDP

common DC precursor

cMoP

common monocyte progenitor

DCs

dendritic cells

E. coli

Escherichia coli

E9·5

embryonic day 9·5

EAE

experimental autoimmune encephalomyelitis

EMP

erythromyeloid progenitors

G‐CSF

granulocyte colony‐stimulating factor

GMPs

granulocyte and macrophage progenitors

GMPs

granulocyte–macrophage progenitors

IRF8

interferon regulatory factor 8

LPS

lipopolysaccharide

MDP

monocyte and DC progenitor

MPS

mononuclear phagocyte system

NETs

neutrophil extracellular traps

pre‐cDC

cDC precursor

RES

reticuloendothelial system

SatM

segregated nucleus‐containing atypical Ly6C^lo monocyte

VEGF‐α

vascular endothelial growth factor α

WHIM

warts, hypogammaglobulinaemia, immunodeficiency and myelokathexis

YSMP

yolk sac‐derived myeloid‐based progenitor

Introduction

Under healthy physiological conditions, the human body maintains a steady number of leucocytes by means of cell proliferation, differentiation, survival and cell death. The bone marrow is regarded as the primary haematopoietic factory for the production of the majority of adult immune cells. Following their egression into the circulation and tissues, these cells will fulfil their purpose and then eventually die, only to be replaced by newly produced younger cells thought to keep the immune system alert at all times. The importance of this fine balance can be appreciated in diseases such as haematological cancers where a substantial number of abnormal immune cells are present or, conversely, where a deficit of specific immune cell subsets consequently results in increased susceptibility to opportunistic infections. ¹ , ² , ³ , ⁴

During inflammation, this finely balanced sequence must quickly adapt to produce sufficient immune cells in order to combat injury or infection and replace those dysregulated by the inflammatory challenge. The cellular kinetics and regulation of the inflammatory response encompass several cell types including sensory C‐fibres, white blood cells, endothelial cells and fibroblasts. Here, we focus on the cellular kinetics of professional phagocytes, a group of innate myeloid immune cells (namely monocytes, dendritic cells, macrophages and neutrophils) specialized, but not limited to, their ability to phagocytose foreign bodies. Tissue macrophages are sentinels that reside within every tissue in an organism and act as first responders to an infectious agent. Depending on the tissue, macrophages are initially derived from progenitors within the yolk sac or fetal liver ⁵ , ⁶ , ⁷ , ⁸ and play an important role during tissue development and maintenance. ⁶ , ⁹ , ¹⁰ , ¹¹ , ¹² , ¹³ , ¹⁴ On the other hand, neutrophils are recognized for their mechanisms involved in pathogen clearance (e.g. reactive oxygen species production, degranulation of antimicrobial proteins, neutrophil extracellular traps (NETs)). Patients diagnosed with neutropenia are often more susceptible to infections, ¹⁵ , ¹⁶ which highlights the need for an adequate production of these cells. Monocytes also aid in the phagocytosis of pathogens and apoptotic cells, whereas dendritic cells (DCs) activate the adaptive immune response by migrating from the periphery to draining lymph nodes. The absence of monocytes and DC has been observed in patients bearing mutations in the transcription factor, interferon regulatory factor 8 (IRF8), as a result, these individuals are more predisposed to infections. ¹

Inflammation is necessary to protect an organism from infectious agents, yet an overactive immune response can result in further tissue damage. This can be observed in chronic inflammatory diseases such as rheumatoid arthritis where a constant recruitment of monocytes to the synovial tissue has been observed in humans. ¹⁷ Therefore, though inflammation has been historically recognized as a salutary reaction, ¹⁸ ‘more is more’ is not often the case. Striking the fine balance of an essential yet limited inflammatory response may present as a potential therapeutic opportunity, highlighting the importance of understanding the kinetics of immune cells in health and disease.

Neutrophil kinetics

Neutrophils are an indispensable component of the innate immune system, if not the most important. Within the circulation, neutrophils constitute the largest proportion of human circulating leucocytes and estimated to be found at ~ 4·2 x10⁹ cells per litre of blood although this can vary with age, ethnicity and sex. ¹⁹ , ²⁰ Neutrophils are often the first white blood cell recruited to sites of injury, which is facilitated by both the upregulation and expression of membrane adhesion molecules on the endothelium and neutrophils. ²¹ , ²² Increased levels of neutrophil chemotactic factors such as IL‐8 (CXCL8) have been observed early on during models of human inflammation, which aids in the guidance of neutrophils to the site of inflammation. ²³ , ²⁴ The initial recognition of pathogens by tissue‐resident macrophages is also partly responsible for the recruitment of neutrophils ²⁵ although other resident cells can also modulate their infiltration. ²⁶ Recent studies in mice and humans have reported a role for vascular endothelial growth factor α (VEGF‐α) released from cDC1 in neutrophil recruitment in cutaneous infections. ²⁷

The generation of neutrophils involves a series of maturation steps starting with granulocyte–macrophage progenitors (GMPs) in the bone marrow. These cells give rise to pre‐neutrophils, which subsequently develop into immature (band) and mature (segmented) neutrophils, which egress into the circulation. ²⁸ , ²⁹ , ³⁰ CXCR4 is a key chemokine receptor involved in the retention of neutrophils within the bone marrow via interaction with CXCL12, whereas CXCR2 activation promotes the mobilization of neutrophils from the bone marrow into the blood. ³¹ , ³² , ³³ The importance of the CXCR4 axis in the regulation of neutrophil egression can be highlighted in warts, hypogammaglobulinaemia, immunodeficiency and myelokathexis (WHIM) syndrome patients who primarily suffer from an autosomal, dominant, gain‐of‐function mutation in CXCR4. ¹⁵ , ¹⁶ Consequently, these patients suffer from defected neutrophil egression and thus circulating neutropenia, making them more vulnerable to infections.

Deuterium labelling acts in a non‐cytotoxic manner to label dividing cells in vivo. ³⁴ , ³⁵ Using this approach to monitor human neutrophil development, following the last proliferation in the bone marrow, the mean transit time before neutrophils enter the circulation has been estimated at 5·8 days. ¹⁹ Once within the circulation, these cells have an incredibly short half‐life of approximately 19 hours. The short half‐life of these cells may reflect their function. Whilst homeostatic functions have been assigned to neutrophils, ³⁶ , ³⁷ these cells are renowned for their defensive mechanisms against invading pathogens such as the production of reactive oxygen species, antimicrobial peptides and NETosis. ³⁸ , ³⁹ These mechanisms are not pathogen‐specific and can therefore result in collateral damage to host tissues, and consequently, it is thought these cells are likely programmed for a quick cell death in order to prevent excessive damage.

Once in the circulation, neutrophils can either be found within the circulating pool where they are readily accessible for blood sampling or be located in a marginating pool. Following stimuli including infection, trauma, the administration of adrenaline or an intense burst of exercise, this marginated pool can be mobilized into the circulating pool in humans. ⁴⁰ , ⁴¹ It has been calculated that the marginating pool makes up approximately 50% of the total neutrophil pool. ¹⁹ Neutrophil transit within tissues refers to how quickly cells pass through the capillary beds. Within the spleen and bone marrow, it takes approximately 10 minutes for neutrophils to transit. ⁴² , ⁴³ Therefore, the period of time cells spend within tissues may factor in determining the marginated fraction.

Neutrophilia is commonly associated with bacterial infections but can arise from other stimuli. In human models of local and systemic inflammation, neutrophilia accounts for the increase in the total white blood cell count at early time‐points. ²³ , ⁴⁴ , ⁴⁵ , ⁴⁶ , ⁴⁷ Of note, left shift refers to the presence of an increased amount of immature neutrophils within the circulation. ⁴⁸ This observation has been associated with cancer progression ²⁸ and has more recently been observed in severe COVID‐19 patients, where these cells exhibit an immunosuppressive profile. ⁴⁹ Elevated levels of systemic granulocyte colony‐stimulating factor (G‐CSF) during inflammation can lead to both the downregulation of CXCR4 ⁵⁰ and the increased levels of CXCR2 ligands, ⁵¹ which likely leads to an increase in neutrophil egression. Additionally, an increase in the number of neutrophil progenitors during inflammation can also contribute to the elevation of circulating neutrophils, ²⁸ in addition to neutrophils from the marginating pool. During the resolution of inflammation, neutrophils are eventually cleared from tissues to restore tissue homeostasis. Neutrophil death can occur in various ways including apoptosis, necrosis, NETosis and autophagy. Efferocytosis refers to the phagocytosis of apoptotic cell bodies. The recognition of ‘eat me’ signals such as phospholipid phosphatidylserine on apoptotic cells facilitates the recognition by macrophages and monocyte‐derived cells to efferocytose apoptotic bodies. ¹⁴ , ⁵² Under steady physiological conditions, senescent neutrophils can also home back to bone marrow by upregulating CXCR4, where they will also be efferocytosed by resident macrophages. ⁵³ , ⁵⁴

These data demonstrate a significant amount of knowledge regarding neutrophils kinetics in both steady state and inflammation. The next step is to truly understand the kinetic changes within each compartment of the body, that is bone marrow, blood and tissue, in addition to the cues that skew the lifespan of these cells during pathology, which may allow for future targeting to promote an adequate response.

Macrophage kinetics

Tissue‐resident macrophages form a network of cells that reside throughout the organism. Macrophage nomenclature has evolved and is based on anatomical location or after the scientist who discovered the macrophage population, for example microglia in the brain, osteoclasts in the bone or Kupffer cells in the liver and Langerhans cells in the epidermis.

The idea that all macrophages arise from monocytes stemmed from studies as early as 1926 demonstrating the ability of monocytes to develop a macrophage phenotype both in vitro ⁵⁵ and in vivo. ⁵⁶ In 1969, Ralph van Furth and colleagues proposed the reticuloendothelial system (RES) was too broad a definition and coined the term the ‘Mononuclear Phagocyte System’ (MPS) including monocytes and macrophages. Furthermore, Van Furth and colleagues separated macrophages into two clear categories: ‘free’ or ‘fixed’ macrophages. When distinguishing the macrophages of the spleen, lymph nodes and bone marrow, ‘free’ macrophages were described as monocyte in origin, while the origin of the ‘fixed’ macrophages in these organs remained undetermined. ⁵⁷ This insight into a division of labour of macrophage ontogeny is reflected at this time with the observation that macrophage development was noted within the yolk sac prior to bone marrow haematopoiesis. ⁵⁸ , ⁵⁹ These yolk sac macrophages appeared one day earlier than monocytes in mice and was one of the earliest observations – to the best of our knowledge – to report macrophage development independent of monocytes. ⁵⁹ Furthermore, investigations into the kinetics of monocytes and macrophages by Ralph van Furth and Zan Cohen, using radioactive thymidine, demonstrated low labelling in peritoneal macrophages compared with blood monocytes. ⁶⁰ Unaware at the time, these data supported the maintenance of macrophage populations independently of monocytes. More recently, the use of genetic reporter and fate‐mapping techniques has confirmed that tissue‐resident macrophages are embryonically (yolk sac and/or fetal liver) derived and can be maintained with little or significant monocyte input depending on the tissue of concern ⁵ , ⁶ , ⁷ , ⁸ , ⁶¹ , ⁶² , ⁶³ (Figure 1).

Figure 1.

Tissue macrophage kinetics with age. Illustrative diagram demonstrating the relative and differential rates of blood monocyte contribution to tissue macrophage populations. Adapted from Liu et al. (2019).

Within the developing embryo, haematopoiesis occurs in a sequential manner, initially occurring within the yolk sac (primitive haematopoiesis) before transferring to the fetal liver until birth where haematopoiesis is ultimately transferred to the bone marrow (definitive haematopoiesis). ⁶⁴ Primitive haematopoiesis begins in the yolk sac, where erythromyeloid progenitors (EMPs) give rise to erythrocytes, macrophages and mast cells. ⁶ , ⁶⁵ A similar yolk sac‐derived myeloid‐based progenitor (YSMP) has been identified in humans, however lacks an erythroid gene signature. ⁶⁶ Downstream of the EMP, a pre‐macrophage precursor has been identified in mice, which arises in the yolk sac before colonizing embryonic tissues around embryonic day 9·5 (E9·5) at the same time as organogenesis. ⁶ These pre‐macrophages are subjected to tissue‐specific signals, which help sculpt a tissue‐specific resident macrophage phenotype. ⁶⁷ Recent studies on macrophage development in rodents have been corroborated in humans where an early monocyte‐independent primitive wave and later fetal liver monocyte‐derived wave have been described. ⁶⁶

Following birth, resident macrophages can persist throughout life with or without monocyte input depending on the tissue compartment under steady state ⁸ , ⁶² , ⁶³ (Figure 1). More recently, a novel fate‐mapping model utilizing the Ms4a3 gene demonstrated in a more specific quantitative manner the monocyte contribution to several tissue macrophage compartments. ⁶³ In tissues such as the brain, skin and liver, monocytes do not replace the embryonically derived microglia, Langerhans cells nor Kupffer cells. Whilst this has been clearly demonstrated in mice, further clarity is required within the human setting. Interestingly, Langerhans cells of the skin have been shown to persist of donor origin – ten years following human hand allograft. ⁶⁸ However, others have found that following bone marrow transplantation, the majority of Langerhans cells were donor‐derived within 3 months. ⁶⁹ , ⁷⁰ , ⁷¹ It is important to note chemotherapeutic or other pharmacological regimes could impact on these studies. The ontogeny of alveolar macrophages in humans has also been examined in humans where contrasting findings have again been observed. ⁷² , ⁷³ , ⁷⁴ An interesting study by Réu and colleagues took advantage of atmospheric ¹⁴C to estimate the turnover rate of human microglia. ⁷⁵ Increases in atmospheric ¹⁴C in the 1950s due to nuclear bomb testing resulted in the increased presence of ¹⁴C within the atmosphere and subsequently DNA of newly formed cells, which in turn could provide insight into the age of a cell. ⁷⁶ Réu and colleagues demonstrated that human microglia have a lifespan of approximately 4·2 years and renew at a rate of 28% per year. ⁷⁵ Table 1 lists examples of studies where the longevity of macrophage populations has been examined in both murine models and the human setting.

Table 1.

Longevity of phagocytes. Summary of longevity of phagocyte populations in mouse and humans. Lifespans and half‐lives have been stated where available. TBC, to be confirmed

	Human	Mouse
Blood neutrophil	Half‐life ~ 13–19 hours 19	Half‐life ~ 12·5 hours 184
Blood classical monocytes	Lifespan ~ 1·0 days 106	Half‐life ~ 20 hours 8
Blood intermediate monocytes	Lifespan ~ 4·3 days 106	TBC
Blood non‐classical monocytes	Lifespan ~ 7·4 days 106	Half‐life ~ 2·2 days 8
Blood DC	TBC	TBC
Langerhans cell	Donor‐derived cells from hand allograft transplant maintained up to 10 years 68 Langerhans cells are donor‐derived following bone marrow transplant after 3 months 69 , 70	Following bone marrow transplant, Langerhans cells remained of recipient origin up to 18 months 185
Microglia	Lifespan ~ 4·2 years and median renewal rate of 28%/year 75	Estimated ~ 96 days for entire microglia population to self‐renew 186
Alveolar macrophage	9·4 years post‐lung transplant ~ 73% of alveolar macrophage are recipient‐derived 73 3·5 years post‐lung transplantation > 87% of alveolar macrophages remain donor‐derived 72	8 months following bone marrow transplantation, negligible donor‐derived alveolar macrophages observed (1–5%) 187
Liver mononuclear phagocyte	11 years post‐liver transplant, donor‐derived CD14+ CD16‐ MNP observed 188	TBC
Testicular macrophages	TBC	Long‐lived macrophages with slow turnover measured over 24 weeks 117
Gut macrophages	Macrophages isolated from the duodenum–proximal jejunum are all recipient‐derived by 52 weeks following duodenal transplant 189	Long‐lived macrophages observed up to 35 weeks within the submucosa and muscularis of ileum 190
Cardiac macrophages	8·8 years following heart transplant, <1% of CCR2‐ macrophages are recipient‐derived 191	Cardiac macrophages gradually replaced with age 192

In the absence of tissue macrophages, a reduced infiltration of granulocytes has been observed during inflammation, ²⁵ highlighting macrophages as one of the many cell types involved in the initiation of the immune response. Following tissue injury of infection, an interesting phenomenon known as the ‘macrophage disappearance reaction’ occurs, first described in 1963. ⁷⁷ , ⁷⁸ This observation has been noted by several groups during experimental peritonitis, where a reduced recovery of resident macrophages was reported. ⁷⁹ , ⁸⁰ , ⁸¹ This has also been extended to other tissues, such as the lung where a low number of alveolar macrophages are recovered following influenza challenge in mice ⁸² and the liver where a lower number of Kupffer cells are present during inflammation. ⁸³ , ⁸⁴ Zhang and colleagues recently showed that macrophages form clots and adhere to tissues accounting for the reduced recovery of these cells, which could be reversed by the use of anticoagulants. ⁸¹ Following the resolution of inflammation, the recovery of resident macrophage numbers may occur by proliferation, ⁸⁵ repopulation by monocyte‐derived cells ⁸ , ⁶³ , ⁷⁹ , ⁸⁶ , ⁸⁷ , ⁸⁸ or a combination of both. ⁸⁹ , ⁹⁰ As expected, exceptions to the macrophage disappearance paradigm have been observed in T helper cell type 2 immune response, where tissue‐resident macrophages proliferate to combat infection rather than depend on monocyte recruitment. ⁹¹

Monocyte kinetics

In contrast to macrophages, monocytes and DC are derived from bone marrow progenitors. Initially, the monocyte and DC progenitor (MDP) was demonstrated to lack neutrophil potential yet give rise to monocytes via the common monocyte progenitor (cMoP). ⁹² , ⁹³ , ⁹⁴ Although more recently, the cMoP has been proposed to descend from the granulocyte and macrophage progenitors (GMPs) bypassing the MDP stage in both mice ⁶³ and humans. ⁹⁵ In this study, the GMP and MDP were suggested to generate monocytes through two pathways, a GMP → cMoP → monocyte pathway and a MDP → monocyte pathway that lack a cMoP intermediate stage. ⁶³ Commitment to monocyte development at the cMoP stage is dependent on the transcription factor IRF8. ⁹⁶ This is consistent with the observation in patients bearing mutations in IRF8 who are also deficient of circulating monocytes. ¹ Recently, a proliferative CXCR4^hi CCR2^lo transitional pre‐monocyte population was described in mice and humans within the bone marrow, which eventually downregulates CXCR4 and upregulates CCR2, resulting in the egression of mature classical monocytes into the circulation. ⁹⁷ , ⁹⁸ , ⁹⁹

The kinetic profiles of circulating monocyte subsets have been examined in mice with the use of BrdU, where a sequential appearance of labelled classical monocytes and then non‐classical monocytes appears in the circulation. ⁸ This is owed to the fact that Ly6C^hi classical monocytes convert into Ly6C^lo non‐classical monocytes. ⁸ , ⁶³ , ⁹⁴ , ¹⁰⁰ , ¹⁰¹ The half‐life of classical and non‐classical monocytes in mice is estimated at 20 hours and 2·2 days, respectively.

Early insights into the kinetics of human monocytes stemmed from studies nearly fifty years ago with the use of tritiated thymidine where the average monocyte lifespan was estimated at 4·25 days. ¹⁰² Deuterated glucose labelling studies proposed a half‐life of 2·2 days for CD14⁺ classical monocytes. ¹⁰³ With regard to monocyte subsets, studies have demonstrated differences in the kinetic profiles of circulating CD14⁺ and CD16⁺ monocytes in patients following haematopoietic stem cell transplantation. ¹⁰⁴ Using deuterium labelling, a sequential appearance of monocyte subsets has been observed within the circulation. ¹⁰⁵ , ¹⁰⁶ Using mathematical modelling, it was estimated classical monocytes are retained within the bone marrow for approximately 1·6–2 days between the last mitotic division and entry into the circulation ¹⁰⁵ , ¹⁰⁶ (Figure 2). These cells then circulate for approximately 1 day, whereas non‐classical monocytes circulate for a longer period of time (~7·5 days). ¹⁰⁶ These results are akin to those observed in mice, ⁸ rats ¹⁰⁷ and macaques ¹⁰⁸ , ¹⁰⁹ most likely due to a conserved developmental relationship between monocyte subsets.

Figure 2.

Overview of monocyte kinetics in health and inflammation. Under steady physiological conditions, classical monocytes arise from cMoP and reside within the bone marrow for approximately 1·6 days before being released into the circulation. Once within the circulation, these cells mature into intermediate and then non‐classical monocytes, which circulate for 4 and 7 days, respectively. During systemic inflammation, classical monocytes are rapidly released into the circulation. Following local injury, classical monocytes are initially recruited later followed an intermediate monocyte phenotype, although the origin of the cells is unknown (dashed lines). HSC, haematopoietic stem cell; cMoP, common monocyte progenitor.

Non‐classical monocytes reside within the circulation for a longer period of time possibly due to their function within the circulation where they are constantly monitoring the endothelium for damage. ¹¹⁰ , ¹¹¹ These cells represent a terminally differentiated monocyte and may consequently even represent a ‘blood macrophage’. ⁸ Supplemental components encourage non‐classical monocyte survival include CX₃CL1 ¹¹² and TNF. ¹¹³ On the other hand, classical monocytes are continuously recruited to repopulate tissue mononuclear phagocyte compartments, ⁶¹ , ⁶³ , ¹⁰⁴ , ¹¹⁴ , ¹¹⁵ , ¹¹⁶ , ¹¹⁷ which may explain their shorter circulating lifespan. Liu et al. demonstrate the rate at which monocytes replace tissue macrophages in mice is tissue‐specific, for example kidney macrophages are gradually replaced overtime but to a lesser extent than lung alveolar macrophages or dermal macrophages ⁶³ (Figure 1). The determinants of these rates remain unknown, although the macrophage niche theory proposes a niche is filled by the competition between an embryonic macrophage and a monocyte‐derived cell. ¹¹⁸ Therefore, the differences between tissue microenvironments most likely dictate the rate of macrophage replacement.

In an inflammatory setting, a reduction in the number of circulating monocytes has been observed in both mice and humans, hours following intravenous endotoxin challenge. ⁴⁴ , ⁹⁷ , ¹⁰⁶ , ¹¹⁹ , ¹²⁰ The lung is a primary location where activated monocytes can marginate to following challenge. ⁹⁷ , ¹²¹ Given the narrow diameter of lung capillaries, ¹²² changes in the morphology and size of monocytes following activation may result in hindrances and consequently an increased transit time. However, active retention of monocytes within the lung via CXCR4 retention has also been documented in mice. ⁹⁷ Targeting margination in the lung may be of clinical relevance as this accumulation of monocytes can promote further lung injury. ¹²¹ Following temporary monocytopenia, monocytes can return from various sources. In mice and humans, the spleen has been implicated as a monocyte reservoir, which is deployed in response to specific inflammatory cues. ¹²³ , ¹²⁴ We and others have shown that bone marrow ‘immature’ classical monocytes are rapidly released into the circulation in response to bacterial ¹⁰⁶ and sterile inflammation ¹²⁴ (Figure 2). More recently, patients with severe COVID‐19 present with immature classical monocytes and neutrophils within the circulation, ⁴⁹ suggesting this emergency release of monocytes is also apparent in a viral setting.

Intradermal challenges have allowed for the study of immune cell recruitment to the site of infection in response to various stimuli in humans. ²³ , ¹²⁵ , ¹²⁶ In response to UV‐killed Escherichia coli (E. coli), classical monocytes are observed within the skin as early as 8 hours following challenge ²³ (Figure 2). This observation is akin to mice, where Ly6C^hi classical monocytes are typically the subset recruited ¹²⁷ , ¹²⁸ , ¹²⁹ possibly via a CCR2‐dependent manner. ⁹⁹ , ¹²³ , ¹²⁸ , ¹³⁰ At later time‐points, a Ly6C^lo ‘non‐classical’ phenotype is apparent, but it is thought that this is due to in situ conversion rather than a second wave of monocyte recruitment. ¹²⁷ , ¹²⁸ , ¹²⁹ , ¹³¹ , ¹³² Similarly, in humans, CD16 expression increases over time ²³ and may also represent maturation at the site of infection. While it may seem that classical monocytes are the prime effector subset responsible for inflammation and resolution, the use of Nr4a1‐deficient mice ¹³³ , ¹³⁴ has highlighted a role of non‐classical monocytes in various pathologies including tumour metastasis, ¹³⁵ Alzheimer’s disease, ¹³⁶ experimental autoimmune encephalomyelitis (EAE) ¹³⁷ and vascular homeostasis. ¹¹¹ The role of non‐classical monocytes has recently been extensively covered. ¹³⁸ Of note, following the resolution of inflammation, it is thought that tissues return to baseline homeostasis. Whilst this is true at the symptomatic level, immunological processes have been observed to continue in mice where IFN‐γ triggers a second wave of monocyte recruitment creating an immune‐suppressed environment. ¹³⁹ This may be of importance when considering secondary infections; therefore, targeting this second wave of monocytes may be of clinical relevance.

The question arises whether these recruited inflammatory monocytes engraft into the long‐lived macrophage pool. In models of peritonitis, monocyte‐derived cells persist up to 8 weeks, where their phenotype gradually changes into that of resident macrophages. ⁸ , ⁷⁹ Similar observations have been extended to the liver ⁸⁶ , ¹⁴⁰ , ¹⁴¹ , ¹⁴² and lung. ⁸⁷ , ⁸⁸ , ¹⁴³ It is possible that tissue residence could alter the longevity of monocytes although, in mouse models of experimental autoimmune encephalomyelitis, monocyte‐derived cells do not contribute to the resident microglia pool ¹⁴⁴ yet contribute to pathology, ¹⁴⁵ which possibly highlights the microglia as a unique population owed to their unique location within the blood–brain barrier. Whether monocyte‐derived cells exhibit the same function as their resident macrophage counterparts is of key importance. In a mouse model, the engraftment of monocyte‐derived cells into the lung was examined. After ten months, the graft cells showed a very similar transcriptome to alveolar macrophages and only exhibited a difference of 330 differentially expressed genes. ⁸⁸ In a separate infection study, the replacement of alveolar macrophages with monocyte‐derived cells in response to herpesvirus resulted in protection against house dust mite‐induced asthma compared with mice without initial exposure to herpesvirus. ⁸⁷ Similar findings have recently been documented, where initial exposure to influenza resulted in subsequent protection from Streptococcus pneumoniae due to the recruitment and engraftment of monocytes to the alveolar niche. ¹⁴³ These studies demonstrate, in addition to ontogeny, the context in which monocytes are recruited and the type of stimuli may also shape the function of these cells.

Under pathological conditions, monocyte‐like populations have been observed, and YM1⁺ Ly6C^hi monocytes are greatly expanded within the bone marrow, blood and spleen of mice following intravenous lipopolysaccharide (LPS) challenge where these cells exhibit immunoregulatory properties and aid in tissue repair (Ikeda et al., 2018). In the case of fibrosis, a segregated nucleus‐containing atypical Ly6C^lo monocyte (SatM) has been documented, although they do not arise from the MDP differentiation route and do not arise from Ly6C^hi progenitors (Satoh et al., 2017). Inflammation likely skews ‘healthy’ haematopoiesis; therefore, examining the kinetics of these cells under steady conditions will be the initial step and warrants the need to further investigate the development and kinetics of these cells under pathological conditions.

DC kinetics

A common school of thought has been monocytes are the immature precursor cells to macrophages and DC. However, the identification of the common DC precursor (CDP) that gives rise exclusively to pDC and cDC, but not monocytes, ⁹³ , ¹⁴⁶ , ¹⁴⁷ challenged this view and established a DC‐dedicated lineage in both rodents and humans. Prior to the generation of cDC, CDP initially gives rise to a cDC precursor (pre‐cDC), ¹⁴⁶ , ¹⁴⁸ , ¹⁴⁹ , ¹⁵⁰ , ¹⁵¹ , ¹⁵² which can be skewed to pre‐cDC1 or pre‐cDC2 fate depending on the cues present. ⁹⁶ , ¹⁵⁰ , ¹⁵¹ , ¹⁵³ , ¹⁵⁴ , ¹⁵⁵ In humans, pre‐cDC can be found within the circulation where they can further mature into cDC. ¹⁵⁶ , ¹⁵⁷ , ¹⁵⁸ Whilst mouse pDCs are released as mature cells from the bone marrow and are thought to be derived mostly from lymphoid progenitors, ¹⁵⁹ , ¹⁶⁰ , ¹⁶¹ these observations are yet to be confirmed in humans.

DCs are renowned for their ability to stimulate naïve T cells and subsequently bridge the innate and adaptive immune response. Numerous DC subsets have been identified, each interacting with T cells in various ways. DCs are found at smaller numbers in comparison with other cell types, nevertheless a single DC can interact with up to 500 T cells per hour; ¹⁶² therefore, their low abundance should not undermine their functional relevance. Though two major cDC subsets are widely acknowledged (DC1 and DC2), further heterogeneity has recently been identified within the DC lineage. ¹⁵⁶ , ¹⁵⁷ , ¹⁶³ , ¹⁶⁴ , ¹⁶⁵ , ¹⁶⁶ , ¹⁶⁷

BrdU labelling in mice demonstrated a rapid labelling of splenic DC, initially thought to be attributed to the rapid replenishment from circulating DC precursors. ¹⁶⁸ However, splenic DCs are also proliferative; therefore, the labelling is likely to represent a combination of both in situ proliferation and blood derivation. ¹⁶⁹ Parabiosis studies examined the decay of parabiont‐derived DC in the lymphoid and non‐lymphoid organs and demonstrated DCs are cleared within 10–14 days. ¹⁴⁹ , ¹⁶⁹ Taking into consideration, DC replenishment from blood precursors, division and cell death, Liu et al. calculated that lymphoid organ cDCs are replenished at a rate of 4,300 cells per hour. This rapid tissue replenishment is supported by the rapid turnover of blood cDC in macaques, where these cells were labelled prior to circulating monocytes. ¹⁰⁹ In humans, donor dermal DCs have been identified as early as 18 days following allogeneic haematopoietic cell transplantation and by 56 days were 94% donor‐derived. ¹⁷⁰ Similar studies have also demonstrated that human dermal DCs were replaced by donor origin within 40 days. ¹⁷¹ On the contrary, pDCs have a much slower turnover in comparison with cDC in mice ¹⁷² and macaques, ¹⁰⁹ which is possibly owed to a bias towards cDC production over pDC in the bone marrow. ¹⁷³ Furthermore, the lymphoid origins of pDC ¹⁶⁰ , ¹⁶¹ may also account for the differences in kinetics between myeloid‐derived cDCs.

Akin to monocytes, a reduced number of circulating cDC and pDC have been reported during inflammation in both mice ¹⁷⁴ , ¹⁷⁵ , ¹⁷⁶ and humans. ⁴⁷ , ¹⁷⁷ , ¹⁷⁸ , ¹⁷⁹ , ¹⁸⁰ , ¹⁸¹ The fate of these DCs remains unknown, although DC death is thought to be a factor. ¹⁷⁶ An elegant study by Pasquevich and colleagues demonstrated monopoiesis is favoured over DC production following bacterial infection in mice. ¹⁸² Following TLR4‐mediated inflammation, these mice had reduced numbers of CDP but elevated numbers of cMoP. It is possible the body increases the availability of monocytes to combat infection at the expense of DC; however, this leads to an immunosuppressive state. Interestingly, the rescue of DC from cell death ¹⁷⁶ or increasing DC production via FLT3 ligand ¹⁸³ reduced the inflammatory‐induced immunosuppression and improved survival in mice. Similarly in humans, the number of circulating DCs correlates with survival from secondary infections in sepsis patients. ¹⁷⁷ , ¹⁷⁸ Further exploration into DC kinetics in this setting could present a potential therapeutic target. However, given the diversity of dendritic cell subsets, it is first necessary to understand the foundational biology of these cells and their relationship to one another.

Conclusion

Phagocytes play a crucial role in facilitating the immune response, yet little is known about the tightly regulated processes governing the generation, maturation and disappearance of these cells, which allow them to fulfil their functions. Whilst macrophages are considered as a self‐maintaining population, it is clear this is not the case for all tissues. The question arises – what determines the longevity of macrophages? Why are the rates of monocyte replacement variable between tissues? And consequently, what does this mean functionally? Similarly, given the recent expansion of DC diversity in humans, our knowledge regarding the relationship of the cells to one another, in addition to their individual functions, remains limited. This review summarizes both our current understanding and highlights the gaps in our knowledge of one of the foundational aspects of phagocyte biology. By establishing the kinetics and turnover of these cells in addition to the regulatory mechanisms behind, this may allow for therapies to fine‐tune the immune response.

Acknowledgements

AP is supported by Fondation Gustave Roussy (including the Paediatric Campaign). F.G. is supported by Singapore Immunology Network (SIgN) core funding and Singapore National Research Foundation Senior Investigatorship (NRFI) NRF2016NRF‐NRFI001‐02 and is a European Molecular Biology Organization (EMBO) YIP awardee. SY is supported by the Israel Science Foundation Personal Grant (192/20).

Disclosures

No potential competing interest was reported by the authors.

Data availability statement

Data sharing is not applicable to this article as no new data were created or analysed in this study.