FcRn mediates fast recycling of endocytosed albumin and IgG from early macropinosomes in primary macrophages

It has long been appreciated that IgG has a long half-life relative to other serum proteins and, in the 1990s, the so-called neonatal IgG Fc receptor (FcRn) was shown to be expressed throughout adult life and to function as a ‘protection’ receptor that acts by rescuing IgG that had been internalised through endocytosis by endothelial cells from degradation. Subsequently, FcRn was also shown to extend the half-life of serum albumin (Chaudhury et al., 2003). It is now clear that FcRn rescues endocytosed IgG and albumin from lysosomal degradation by capturing these ligands in acidic intracellular compartments, prior to their delivery to lysosomes, and recycling them back to the cell surface, where they dissociate at neutral pH and are released, thereby extending their lifetime within the circulation (Roopenian and Akilesh, 2007; Ward and Ober, 2009). In FcRn-deficient mice, the half-life of IgG and albumin are both reduced to ∼1 day, representing a 4–5-fold reduction for IgG (Roopenian et al., 2003) and a 1.5–2-fold reduction for albumin (Chaudhury et al., 2003). Analysis of clearance rates in conditional FcRn knockout mice demonstrated that endothelial and haematopoietic cells are the major sites of FcRn-mediated IgG homeostasis (Montoyo et al., 2009).

FcRn is a membrane-bound heterodimer consisting of a MHC class I-like heavy chain (∼40 kDa) and a non-covalently associated light chain, β2 microglobulin (β2m) that is common to all MHC class I molecules. FcRn has a classical MHC class I-fold; however, the peptide-binding groove of the α1 and α2 domains is obscured and does not bind peptide ligands (Burmeister et al., 1994a). The IgG-binding site in FcRn involves the α2 and β2m domains of FcRn and pH-dependent salt bridges mediated by two histidine residues of the Fc domain (Burmeister et al., 1994b). The binding site for albumin is on the opposite face of FcRn from the contact sites with Fc and involves a histidine residue (Andersen et al., 2012). FcRn is found in all mammals, although there is species specificity in the interaction with its ligands (Ward and Ober, 2009), a characteristic of the receptor that is relevant to studies of the human ligands in mouse models.

At physiological pH, FcRn does not interact with ligands (Martin et al., 2001). The pH dependence of FcRn binding to ligand defines the intracellular location where the interaction takes place. Albumin is internalised by fluid phase endocytosis, possibly by the efficient macropinocytosis pathway, a pathway highly active in endothelial cells, macrophages and dendritic cells (Lim and Gleeson, 2011; Lim et al., 2015). Cell surface receptors may also be involved in the endocytosis of the FcRn ligands, especially for IgG via Fcγ receptors on haematopoietic cells. Ligands interact with FcRn in the acidic (pH 6.0) endosomal compartments and the complex is recycled to the cell surface where the ligand dissociates from the receptor at neutral pH (Tesar and Björkman, 2010). Analysis of the recycling of IgG and albumin in cell lines, indicates that FcRn recycling occurs via the Rab11 (herein referring to Rab11a and Rab11b forms) recycling endosomes to the plasma membrane (Chia et al., 2018). Live imaging has revealed that GFP-tagged FcRn-positive tubules emanate and break off from endosomal structures, and these tubules are likely to represent transport carriers (Ward et al., 2005). Although these studies using immortalised cells have been informative, the pathways for the uptake of ligand (Anderson, 2014) and recycling, in primary cells is poorly defined. Many studies have investigated internalised IgG by loading the FcRn under acidic conditions, a non-physiological pathway for ligand uptake (Anderson, 2014). Moreover, there have been no reported studies to track the internalisation of albumin by primary cells.

A deeper understanding of the cell biology of ligand uptake pathways in primary cells will assist in a rational approach to the design of recombinant therapeutics that can be recycled via FcRn to extend their half-life (Andersen et al., 2012; Kuo et al., 2010). A key question is whether IgG and albumin share a similar pathway for FcRn–ligand recycling in primary cells or whether there are differences in the uptake and/or recycling pathways. Here, we have analysed the recycling of albumin and IgG in primary mouse macrophages expressing the human FcRn, and have identified a novel fast recycling pathway from early macropinosomes that protects these ligands from degradation.

Macropinocytosis is likely to be the relevant pathway for human serum albumin (HSA) endocytosis in vivo as it represents the most efficient form of fluid phase uptake (Lim and Gleeson, 2011; Swanson, 2008). It is important to analyse macropinocytosis in primary cells as this pathway is considerably more efficient in primary cells compared with immortalised cultured cells. Macrophages are considered a major cell type for the recycling of FcRn ligands in vivo (Akilesh et al., 2007; Montoyo et al., 2009). Here, we used bone marrow-derived macrophages (BMDM) from a mouse line which harbours a knockout allele of the mouse FcRn α-chain (Fcgrt^tm1Dcr) and also expresses the human FcRn α-chain cDNA (FCGRT), termed mFcRn^–/– hFcRn^Tg/Tg cells (Roopenian et al., 2010). Previous studies have shown that the hFcRn expressed in the mFcRn^–/– hFcRn^Tg/Tg mice is functionally active (Petkova et al., 2006; Viuff et al., 2016). This mouse line, mFcRn^–/– hFcRn^Tg/Tg, will be referred to as hFcRn^Tg/Tg mice throughout this study. For comparison, an mFcRn^–/– mouse line, which lacks FcRn, was also used throughout the study. Antibodies to the α-chain of human FcRn (HPA012122) detected a major band of ∼40 kDa in BMDMs lysates from hFcRn^Tg/Tg mice, corresponding to the expected size of the human FcRn heavy chain (Fig. S1). A faint slightly higher Mr band was present in the mFcRn^–/– sample stained with HPA012122 antibodies; however, this faint band was non-specific as it was not detected with another antibody to the human FcRn heavy chain (B-8 antibodies) (data not shown). Immunofluorescence staining of macrophages from hFcRn^Tg/Tg mice showed that hFcRn was predominantly localised to punctate and donut-like structures through the cytoplasm (Fig. S1), whereas no staining was observed in mFcRn^–/– or wild-type BMDMs (Fig. S1). Collectively, these findings indicate that HPA012122 antibodies are specific for hFcRn.

We analysed Alexa Fluor 488-conjugated HSA (HSA–AF488) uptake in BMDMs and observed intense intracellular fluorescence with 100 µg/ml HSA–AF488 after 15 min uptake at 37°C (Fig. S1C). BMDMs were then incubated with both HSA–AF488 and a known marker for uptake into macropinosomes, namely 70 kDa dextran conjugated to TxRed, as 70 kDa dextran is predominantly internalised by macropinocytosis (Kerr et al., 2006; Lim et al., 2012, 2008). After incubation at 37°C for 15 min, HSA–AF488 was detected in the large structures (0.2–2 μm in diameter), a size compatible with the size of macropinosomes, which were also positive for TxRed-conjugated dextran (Fig. 1A), in both hFcRn^Tg/Tg (Fig. 1A) and mFcRn^–/– cell (data not shown). The level of uptake of the HSA–AF488 by hFcRn^Tg/Tg or mFcRn^–/– BMDMs was very similar, as quantified by flow cytometry (Fig. 1D), indicating that the FcRn is not required for initial uptake of the albumin. Moreover, the uptake of HSA–AF488 by hFcRn^Tg/Tg or mFcRn^–/– BMDMs was dramatically reduced (>70% reduction) after treatment with the selective macropinosome inhibitor 5-(N-ethyl-N-isopropyl) amiloride (Koivusalo et al., 2010; West et al., 1989), as assessed by flow cytometry and immunofluorescence, confirming the identity of the large endosomes as macropinosomes (Fig. 1B,C).

Next, we determined the fate of endocytosed HSA in BMDM with and without hFcRn by performing a 15 min pulse with 100 µg/ml HSA–AF488 followed by a chase. Surprisingly, we found that by the end of the 15 min chase, the intracellular HSA–AF488 fluorescence had dramatically waned with very little detected in either hFcRn^Tg/Tg or mFcRn^–/– BMDMs (Fig. 2A, HSA only). To determine whether degradation was contributing to the loss of fluorescence, particularly in the mFcRn^–/– cells, we treated cells with a cocktail of protease inhibitors throughout the pulse and the chase. The fluorescence in the protease inhibitor-treated cells was maintained in the mFcRn^–/– cells, as assessed by confocal microscopy and flow cytometry (Fig. 2A,C,E), indicating that the internalised HSA–AF488 was rapidly degraded in the mFcRn^–/– BMDMs (Fig. 2A,E). To directly analyse the proteolytic degradation of internalised HSA, we used DQ-Red-tagged BSA (DQ–BSA), a self-quenched conjugate of albumin that exhibits fluorescence at 620 nm upon proteolytic degradation in the late endosome/lysosome (Daro et al., 2000; Santambrogio et al., 1999). BMDMs from mFcRn^–/– mice were pulsed with both HSA–AF488 and DQ–BSA for 15 min and then chased at 37°C for 10 min. After the 15 min pulse, HSA–AF488 was detected in large macropinosomes, whereas there was minimum fluorescence at 620 nm (Fig. 3A). After the 10 min chase, there was a very strong signal at 620 nm in endosomal structures dispersed throughout the cytoplasm of CSF-1-treated BMDMs (Fig. 3A). These DQ-Red-positive structures showed extensively colocalisation with HSA–AF488. HSA–AF488 was also detected in very large endosomal structures devoid of 620 nm fluorescence, and these large structures were located in the periphery of the cell (Fig. 3B, arrows). These findings demonstrate, first, that albumin is rapidly degraded in activated BMDMs in the absence of FcRn, and second, that the proteolytic degradation of DQ–BSA occurs downstream of the large early macropinosomes, most likely after subsequent fusion of the maturing macropinosomes with late endosomes (Kerr et al., 2006). Thus, for FcRn to rescue HSA from degradation, the receptor must bind and divert the ligand in the early stages of the macropinosome pathway.

In contrast to mFcRn^–/– BMDM, following endocytosis of HSA–AF488 in hFcRn^Tg/Tg BMDM, the fluorescent signal rapidly disappeared (by 15 min), regardless of the presence or absence of protease inhibitors, as assessed by optical microscopy and flow cytometry (Fig. 2A,C,E). In addition, DQ–BSA internalised by hFcRn^Tg/Tg BMDM showed a minimal signal at 620 nm after a 10 min chase (Fig. 3A); as BSA can interact strongly with the human FcRn (Chaudhury et al., 2003), this finding indicates that BSA is protected from degradation in the presence of FcRn. To determine whether the disappearance of fluorescence was due to FcRn recycling, we incubated hFcRn^Tg/Tg BMDMs with a previously defined FcRn non-binding HSA mutant, rHSA^H464Q-AF488 (Andersen et al., 2012). In contrast to the disappearance of wild-type HSA, this non-binding HSA mutant showed strong intracellular fluorescence after 15 min chase in the presence of protease inhibitors, and regardless of the presence or absence of FcRn (Fig. 2B,D,F), indicating the recycling of wild-type HSA was promoted by FcRn. To assess recycling and secretion of HSA, we analysed cell lysates and the extracellular medium for HSA by immunoblotting. Similar levels of HSA were detected in cell lysates after the pulse, regardless of the presence or absence of FcRn or whether wild-type or mutant HSA was endocytosed (Fig. 4). Elevated levels of HSA, representing 50% of total endocytosed HSA, were detected in the media from hFcRn^Tg/Tg BMDM after the 15 min chase period compared with media from mFcRn^–/– cells (Fig. 4A,B). In addition, only low levels (1–2%) of the non-binding HSA mutant, rHSA^H464Q–AF488, were detected in the medium of hFcRn^Tg/Tg BMDMs after the chase period (Fig. 4C,D). Thus, hFcRn^Tg/Tg BMDMs were able to recycle wild-type HSA, but not the non-binding mutant, rapidly into the medium after endocytosis, whereas mFcRn^–/– BMDMs were not.

We next investigated the mechanism for HSA recycling from macropinosomes. Macropinosomes are known to tubulate extensively during maturation, and the tubules pinch off to become transport carriers (Kerr et al., 2006). The destinations of these macropinosome-derived transport carriers are not well defined, but have been proposed to be involved in recycling. To assess whether HSA entered tubules derived from macropinosomes, HSA–AF488 pulsed BMDMs were fixed under conditions that preserve the tubular structures, namely with 2% PFA. HSA–AF488 was detected in abundant tubules arising from large Alexa Fluor 488-positive macropinosomes in hFcRn^Tg/Tg BMDMs (Fig. 5A). In contrast, very few HSA–AF488-labelled tubules were visible in mFcRn^–/– BMDMs (Fig. 5A) or with the non-binding HSA^H464Q mutant in hFcRn^Tg/Tg BMDM (data not shown). In both cases the labelled HSA–AF488 was restricted to the body of large macropinosomes, indicating that the partitioning of the soluble HSA into tubules is mediated by the membrane FcRn.

The intracellular location of FcRn was also analysed in activated hFcRn^Tg/Tg BMDM. Intracellular staining required fixation conditions which results in the loss of the tubular domains; however, SNX5 provides a marker for newly formed macropinosomes and is also associated with a subdomain that generates the tubular extensions (Kerr et al., 2006). Dual staining for SNX5 and FcRn in hFcRn^Tg/Tg BMDM revealed extensive colocalisation of FcRn with SNX5 in large macropinosome-like structures. Quantitative analysis indicated that >50% of FcRn colocalised with SNX5. EEA1, a marker for early endosomes, is also recruited to early macropinosomes. Approximately 30% of FcRn also colocalised with EEA1, whereas <12% colocalised with Rab11, a marker of recycling endosomes (Fig. 5B). These results demonstrate that FcRn is predominantly present in large macropinosome structures, and therefore FcRn has the potential to mediate binding and recycling from newly formed macro structures.

Next, we analysed the relationship between the localisation of the ligand HSA and the receptor FcRn over a 10 min chase period (Fig. 5D). After a 15 min pulse with HSA–AF488 there was a 50% colocalisation of HSA with FcRn indicating that the ligand and receptor were in close proximity very early after fluid phase uptake. The association remained high throughout the 10 min chase. BMDMs from mFcRn^–/– had no staining with the antibody to FcRn, as expected (Fig. 5C). We then analysed the level of colocalisation of HSA ligand with SNX5, Rab11 and Lamp1 over a 10 min chase to identify the intracellular trafficking pathway for HSA–AF488 (Fig. 5D–F). In hFcRn^Tg/Tg BMDMs, there was a high level of colocalisation of ligand with SNX5, whereas in the absence of hFcRn, the colocalisation with SNX5 was only ∼10% (Fig. 5D). Thus, the presence of the hFcRn resulted in localisation of HSA to SNX5 domains, a relevant finding as SNX5 domains are considered to generate the tubules for cargo recycling. Only small amounts of ligand was associated with either Rab11 or Lamp1 over the 10 min pulse in the presence of hFcRn (Fig. 5E,F), indicating that the majority of ligand did not enter either recycling endosomes or late endosomes/lysosomes. In contrast, in the absence of hFcRn, HSA–AF488 showed only low levels (∼10%) of colocation with SNX5, and high levels of colocalisation with Lamp1. The colocalisation of HSA with the late endosome/lysosome marker in the absence of FcRn increases over the 10 min chase period, indicating delivery of HSA ligand to the lysosomes. Therefore, taken together, in mFcRn^–/– BMDM, HSA–AF488 was restricted to the body of the large endosomes and transported rapidly to lysosomes. In hFcRn^Tg/Tg BMDMs, however, HSA was detected on abundant tubules emerging from large endosomes, and colocalised with FcRn and SNX5 subdomains, whereas very little was directed towards lysosomes.

To directly visualise tubules emerging from early macropinosomes we performed live imaging of BMDMs following a 5 min pulse with HSA–AF488 (Fig. 6). In hFcRn^Tg/Tg BMDM imaging revealed dynamic formation of fluorescently labelled tubules emerging from large structures in the periphery of the cell. These tubular structures were highly mobile and projected towards the cell surface where the fluorescence intensity then waned, indicating delivery of the fluorescent cargo to the plasma membrane (PM) (Movie 1). In contrast, very few HSA–AF488 tubules were observed in mFcRn^–/– BMDM (Fig. 6A; Movie 2). We analysed >10,000 Alexa Fluor 488 tracks in hFcRn^Tg/Tg BMDMs and mFcRn^–/– BMDMs, collected randomly from the total intracellular HSA–AF488 pool and included both endosomal and tubular structures. The Alexa Fluor 488 tracks showed a higher mean displacement and speed in hFcRn^Tg/Tg BMDMs compared with that in mFcRn^–/– BMDMs (Fig. 6C–E). This quantitative analysis data confirms that the HSA–AF488 was associated with more-dynamic structures, which moved across longer distances in hFcRn^Tg/Tg BMDM compared with mFcRn^–/– BMDMs. We then compared the behaviour of the non-binding rHSA^H464Q–AF488 mutant in live BMDMs. Very few rHSA^H464Q–AF488-labelled tubules were visualised in either hFcRn^Tg/Tg BMDM or mFcRn^–/– BMDMs (Fig. 6B, Movies 3 and 4). Moreover, analysis of Alexa Fluor 488 tracks in hFcRn^Tg/Tg BMDMs and mFcRn^–/– BMDMs showed no difference in the displacement and speed of rHSA^H464Q–AF488 (Fig. 6C–E). This data suggests that HSA–AF488 is only recruited into tubules once bound to FcRn. To exclude the unlikely possibility that tubules are dependent on FcRn-bound ligand for their formation, we labelled the membranes of early macropinosomes in mFcRn^–/– BMDMs by use of the PM label CellMask Deep Red, to visualise the membranes of emerging tubules. After PM labelling with CellMask Deep Red and uptake of HSA–AF488, the HSA ligand was observed in CellMask Deep Red-labelled large macropinosomes close to the PM (Fig. S2). Live imaging revealed CellMask Deep Red-labelled tubules emanating from the HSA–AF488-positive macropinosomes (Movie 5, Fig. S2). The CellMask Deep Red-labelled tubules were devoid of HSA-AF488 in the FcRn^–/– BMDM (Fig. S2), demonstrating that the tubules form independently of FcRn and its ligand. Hence, collectively these data demonstrate that HSA–AF488 is recruited into dynamic tubules emerging from early macropinosomes in FcRn-dependent manner.

A question that arises from these findings, is whether IgG follows a similar intracellular pathway to that of albumin in activated macrophages, given that IgG is not only a ligand of FcRn but also for Fcγ receptors, which have a role in delivery of immunoglobulin–antigen complexes to the MHCII loading compartments for antigen presentation. To compare the transport routes of albumin and IgG, BMDMs were pulsed with 100 µg/ml HSA–AF488 and Alexa Fluor 594-conjugated IgG (IgG–AF594) simultaneously for 15 min in the presence and absence of protease inhibitors. As expected, with a 0 min chase, HSA–AF488 was detected in large macropinosomes in BDMDs treated with either carrier only or with protease inhibitors. The majority of the IgG was associated with the cell surface, with only low levels colocalised with internalised HSA–AF488. After the 15 min chase, the HSA–AF488 signal was greatly reduced in both FcRn^–/– BMDMs and hFcRn^Tg/Tg BMDMs whereas the IgG–AF594 persisted, as assessed by confocal microscopy (Fig. 7A) and also by flow cytometry of BMDM pulsed with 100 µg/ml IgG–AF488 alone (Fig. 7B). To assess whether Fcγ was responsible for cell surface binding and persistence, we treated BMDMs with mouse serum to block the surface Fcγ receptors (Fig. S3) and repeated the uptake experiment with both ligands. Mouse IgG binds (and blocks) Fcγ receptors but is unable to interact with human FcRn. At the end of the 15 min pulse (0 min chase), both Alexa Fluor ligands were detected in large macropinosomes and showed extensive overlap (Fig. 7A). After the 15 min chase, very little signal was detected in the absence of protease inhibitors regardless of the presence or absence of FcRn. In the presence of protease inhibitors there remained a strong signal for both ligands in FcRn^–/– BMDM whereas the signals were very weak in hFcRn^Tg/Tg BMDMs (Fig. 7A). Analysis by flow cytometry of BMDMs pulsed with solely IgG–AF488 confirmed these observations; there was a dramatic reduction in signal in hFcRn^Tg/Tg BMDMs even in the presence of protease inhibitors (Fig. 7B). Hence, if the Fcγ is blocked or saturated by unlabelled mouse IgG, the internalisation of monomeric IgG by fluid phase macropinocytosis is more readily observed and the IgG internalised by this pathway follows similar FcRn-dependent trafficking kinetics to albumin.

The physiological pathways for uptake and recycling of FcRn ligands in primary cells are poorly defined (Anderson, 2014). Here, we have shown that the fluid phase uptake of albumin and IgG by primary macrophages is predominantly undertaken by macropinocytosis and have identified a rapid recycling pathway for albumin from newly formed macropinosomes. Our study demonstrates that FcRn mediates the rapid recycling directly from newly formed macropinosomes in activated macrophages because: (1) ∼50% HSA is recycled into the cell culture within 15 min of internalisation; (2) HSA is detected in tubules emerging from early macropinosomes which were then directed towards the PM; (3) in the absence of FcRn, HSA is neither partitioned into tubules nor recycled, but remains within the body of the macropinosome and is subsequently rapidly degraded; (4) the non-FcRn-binding mutant of HSA, rHSA^H464Q–AF488, is rapidly internalised and degraded in hFcRn^Tg/Tg BMDM, and is not recycled into the culture medium and (5) in fixed BMDMs, FcRn is detected in SNX5 subdomains of the newly formed macropinosomes, domains known to give rise to tubular carriers (Kerr et al., 2006). We also demonstrated that monomeric IgG taken up by fluid phase endocytosis follows a similar pathway to that of albumin.

Macropinocytosis is the major pathway for uptake of HSA by primary macrophages, as demonstrated by the colocalisation with the fluid phase marker of macropinocytosis 70 kDa dextran, into very large endocytic structures and the reduction of HSA endocytosis with amiloride, a selective inhibitor of this pathway (Koivusalo et al., 2010). Macropinosomes are generated from actin-mediated ruffling of the PM. Recent studies have identified two distinct pathways of macropinocytosis in macrophages, wherein macropinosomes are derived from peripheral ruffles or derived from dorsal ruffles following receptor activation (Condon et al., 2018; Lim et al., 2015). Dorsal ruffles give rise to larger macropinosomes than peripheral ruffles (Condon et al., 2018; Lim et al., 2015). Our previous studies have shown that the majority of macropinosomes from activated macrophages are derived from these dorsal ruffles (Lim et al., 2015). Newly formed macropinosomes are known to tubulate extensively, a process dependent on SNX5 and other Bar domain proteins (Kerr et al., 2006). These membrane tubules are precursors of transport carriers considered to transport cargoes to other intracellular destinations. Recycling of membrane cargo, endocytosed by macropinocytosis, back to the PM has been previously demonstrated (Bryant et al., 2007), although the relationship between the early tubulation events and cell surface recycling has not been previously demonstrated. The findings in this paper show that newly formed macropinosomes undergo extensive tubulation, a process involving removal of membrane from the early macropinosome, followed by acquisition of proteolytic enzyme activity, presumably by the maturing macropinosome body fusing with late endosomes/lysosome compartments.

In the absence of FcRn, or the inability to bind FcRn, HSA was rapidly degraded. Activated macrophages have been reported previously to be able to very rapidly (within 10 min) degrade endocytosed soluble proteins (Diment and Stahl, 1985). The FcRn-dependent partitioning of HSA into tubules of the newly formed macropinosomes provides a mechanism to rapidly remove the ligand from the maturing macropinosome body and protect it from proteolytic degradation. The use of self-quenched DQ–BSA showed that newly formed macropinosomes did not proteolytically digest albumin; however, within 10 min of chase following internalisation of DQ–BSA by FcRn^–/– BMDMs, the self-quenched DQ–BSA showed fluorescence at 620 nm, indicating that the soluble conjugate was exposed to a degradative environment. Our findings are consistent with previous studies, which showed that BSA is rapidly degraded by macrophages after internalisation (Diment and Stahl, 1985). Hence, to be protected from degradation, cargo must be removed rapidly from the newly formed macropinosome. The tubules emanating from newly formed macropinosomes allow the receptor and cargos to be returned to the PM instead of being subject to proteolytic digestion. Indeed, this is the first example in which we are aware of a receptor which is directly recycled from newly formed macropinosomes back to the PM. A model summarising our findings is shown in Fig. 8.

What is the source of the FcRn in early macropinosomes? Given the very rapid recycling of albumin, it is likely that hFcRn is internalised from the cell surface via membrane ruffles, as proposed in the model depicted in Fig. 8. An acidic environment is required for binding of FcRn to ligand. Following their biogenesis in macrophages, macropinosomes are known to shrink and acidify (Racoosin and Swanson, 1989; Swanson and Watts, 1995). It was recently reported that the WASH complex regulates V-ATPase trafficking on macropinosomes as an early event in macropinosome maturation in Dictyostelium (Buckley et al., 2016). Hence, it is possible a similar process occurs in mammalian macrophages to acidify early macropinosomes. Also of note is that very little Rab11 colocalised with albumin during recycling in these activated macrophages, indicating that the recycling pathway is independent of Rab11. This finding is in contrast to what is seen in cells that do not have a high macropinocytic activity, where recycling of FcRn-bound ligand occurs via Rab11 recycling endosomes (Chia et al., 2018). Sorting signals have been identified in the cytoplasmic tail of FcRn that are required for transport of FcRn from early endosomes to recycling endosomes (Mahmoud et al., 2017). An interesting question concerns the mechanism for sorting of FcRn into the tubular membranes of the early macropinosomes. One possibility is that this could involve similar sorting machinery to that used in the early endosome tubulation process (Cullen and Steinberg, 2018). However, of note is that Rab4, which is considered to regulate rapid recycling back from early endosomes back to the PM (Maxfield and McGraw, 2004; van der Sluijs et al., 1992), does not colocalise with internalised HSA in early macropinosomes (W.H.T. and P.A.G., unpublished data), indicating that the transport route is Rab4 independent. Other possible candidates for the recycling machinery could include SNX17 (McNally et al., 2017; van Kerkhof et al., 2005), SNX27 (Steinberg et al., 2013), and the retromer (Cullen and Steinberg, 2018) and retriever complexes (McNally et al., 2017). The challenge will be to now define the trafficking machinery involved in this transport pathway in primary macrophages.

Our results also provide a basis for understanding the intracellular itinerary and fate of IgG endocytosed by antigen-presenting cells, depending on whether IgG is monomeric or exists in an immune complex. IgG is not only a ligand of FcRn but also for Fcγ receptors, which have a role in delivery of immunoglobulin–antigen complexes to the MHCII-loading compartments for antigen presentation (Guilliams et al., 2014; Molfetta et al., 2014). Given the concentration of IgG in the circulation, the bulk of the monomeric IgG that enters macrophages will be via macropinocytosis and, based on our findings in this paper, will be recycled by the same FcRn mechanism as albumin. Our data also demonstrated that monomeric IgG bound to surface Fcγ receptors has a long residency time at the PM. On the other hand, immune complexes bind to Fcγ receptors, induce rapid uptake via clathrin-mediated endocytosis (Molfetta et al., 2014) and are delivered to lysosomes, hence avoiding the rapid transport pathway mediated by FcRn in early macropinosomes (Fig. 8B). Internalised complexes may be exposed to FcRn in early or late endosomes and an interaction in these compartments cannot be excluded. Nonetheless, the model proposed in Fig. 8B allows for immune complexes to be delivered to late endosomes and the antigen processing compartment independently of the rapid FcRn recycling pathway identified in this paper. Thus, even though monomeric IgG and IgG–antigen complexes are both potential ligands for FcRn, they are internalised by distinct mechanisms, which in turn determines their intracellular transport fate.

Overall our studies demonstrate the importance of defining FcRn recycling pathways in primary cells. The characteristics of recycling of both albumin and IgG could be relevant for the design of recombinant protein therapeutics that seek to exploit the FcRn pathway for potential half-life extension, notwithstanding that the properties of any particular recombinant protein therapeutic will need to be determined.

Rabbit polyclonal antibodies to FcRn (#HPA012122; 1:200) and protease inhibitors (P1860) were purchased from Sigma-Aldrich. Mouse monoclonal anti-human FcRn (B-8, sc-271745; 1:200) was purchased from Santa Cruz Biotechnology Inc. Goat anti-HSA (#EHALB; 1:600) was purchased from Thermo Fisher Scientific. Lysine-fixable TxRed-conjugated 70 kDa dextran was purchased from Invitrogen (Thermo Fisher Scientific). Rat anti-mouse Lamp1 (#553792, Clone 1D4B; 1:600) was purchased from BD Transduction Laboratories. Rabbit polyclonal anti-Rab11 (#5589; 1:200) and anti-EEA1 (#2411; 1:200) were purchased from Cell Signaling Technology. Mouse antibodies to α-tubulin (Clone 236-10501; 1:5000), DQ-Red BSA (#D12051) and DAPI were purchased from Life Technologies. mCSF-1 (GF053) was purchased from Merck Millipore. Lab-generated rabbit anti-mouse SNX5 antibody has been described previously and was used at 1:1000 (Lim et al., 2012). Secondary antibodies used for immunofluorescence were goat anti-rabbit-IgG conjugated to Alexa Fluor 568 nm, goat anti-rabbit-IgG conjugated to Alexa Fluor 488 nm, goat anti-mouse-IgG conjugated to Alexa Fluor 568 nm and goat anti-mouse-IgG conjugated to Alexa Fluor 488 nm were purchased from Life Technologies. Horseradish peroxidase (HRP)-conjugated sheep anti-rabbit-Ig, anti-mouse-Ig and donkey anti-goat-Ig were purchased from DAKO Corporation (Carpinteria, CA). Whole-mouse serum was obtained from Justine Mintern (University of Melbourne, Bio21 Institute, Australia). CellMask Deep Red was purchased from Invitrogen (Thermo Fisher Scientific; C10046) and used at 1:1000 dilution in Leibovitz's medium.

Plasma-derived human serum albumin (HSA) and purified recombinant IgG₁ (BM4) were labelled with Alexa Fluor 488 (AF488) NHS ester (succinimidyl ester) (Life Technologies, A-20000) or Alexa Fluor 594 (AF594) NHS ester (succinimidyl ester) (Life Technologies, A-37572) respectively, according to the manufacturer's protocol. The recombinant FcRn non-binding albumin variant HSA^H464Q was generated and labelled with Alexa Fluor 488 (AF488) NHS ester (succinimidyl ester) (Life Technologies, A-20000) (rHSA^H464Q–AF488).

FcRn^−/− mice, which harbour a knockout allele of the mouse FcRn α-chain (Fcgrt^tm1Dcr), and FcRn^–/– hFcRn (line 276) Tg mice (hFcRn^Tg/Tg), which have the null mutation for the mouse FcRn gene and a transgene expressing the human FcRn α-chain under the control of the β-actin promoter (Chaudhury et al., 2003), were purchased from Jackson Laboratory (Bar Harbour, ME) and mouse colonies established and maintained under specific pathogen-free conditions in the animal facility of Bio21 Institute, University of Melbourne. All experiments carried out on animals were approved by the Institutional Animal Care and Use Committee.

mFcRn^−/− and hFcRn^Tg/Tg mice (8 to 10 weeks of age) were killed by CO₂ asphyxiation, and BMDMs were generated as previously described (Lim et al., 2012). Cells were seeded in eight-well removable chamber slides (Ibidi, Germany) at 0.3×10⁵ cells/well.

BMDMs were initially starved in C-RPMI [Roswell Park Memorial Institute media (RPMI) supplemented with 10% (v/v) fetal calf serum (FCS), 2 nM glutamine and 100 units/μl penicillin and 0.1% streptomycin (complete RPMI)] overnight in a humidified 10% CO₂ incubator at 37°C. On the next day, monolayers were incubated with either 0.01% DMSO (carrier control) or protease inhibitors in C-RPMI for 4.5 h at 37°C prior to the pulse-chase experiment. The protease inhibitor cocktail (Sigma Aldrich, P1860) contains apotinin, bestatin, E-64, luepeptin and pepstatin A. After 4.5 h, monolayers were washed with PBS buffer three times and pulsed with either HSA–AF488, rHSA^H464Q–AF488, IgG–AF594 or DQ–BSA (100 µg/ml) diluted in C-RPMI supplemented with mCSF-1 (50 ng/ml) in the presence of either 0.01% DMSO or protease inhibitors for 15 min at 37°C. Cell monolayers were then washed 6 times with PBS and chased for 0 min and 15 min in C-RPMI in the presence of either 0.01% DMSO carrier or protease inhibitors.

To quantify HSA recycling, BMDMs grown in six-well plates, were pulsed for 15 min with ligand as above, washed six times with PBS and fresh medium (500 μl) added to the cell monolayers. Conditioned medium was collected after the chase period and centrifuged at 21,130 g for 1 min to remove any cell debris. For cell lysates, monolayers were washed in PBS and lysed in 100 μl of RIPA buffer. 5% of the conditioned medium and 10% of cell lysate from each sample were analysed by immunoblotting. The levels of HSA in the conditioned medium were normalised to the input of cell lysates. The percentage of HSA recycled was calculated by dividing levels of normalised HSA in the conditioned medium by the total HSA levels in the cell lysates at the beginning of the chase period (0 min+protease inhibitors condition).

Monolayers grown on coverslips were fixed in 4% paraformaldehyde (PFA) for 15 min at room temperature, followed by quenching in 50 mM NH₄Cl in PBS for 10 min at room temperature. Cells were permeablised with 0.1% Triton X-100 in PBS for 4 min and blocked in blocking solution (5% FCS and 0.02% sodium azide in PBS) for at least 30 min to reduce unspecific binding. Cells were incubated with primary antibodies diluted in blocking solution for 1–2 h at room temperature and washed six times in PBS. Diluted fluorochrome-conjugated secondary antibodies were added to monolayers and incubated for 30 min at room temperature and then washed six times in PBS. Coverslips were washed in milli-Q water before mounting in Mowoil [10% (w/v) Hopval 5-88, 25% (w/v) glycerol and 0.1 M Tris in milli-Q water]. Confocal microscopy was performed using a Leica SP8 system. Images were collected sequentially for multicolour imaging using a 63×/1.4 NA HCX PL APO CS oil immersion objective. GFP and Alexa Fluor 488 were excited with the 488-nm line of an argon laser, Alexa Fluor 568 and 594 were excited with a 543-nm HeNe laser, Alexa Fluor 647 was excited with a 633-nm HeNe laser, and DAPI with a 405-nm UV laser. Images were collected with pixel dimensions of at least 512×512. For multicolour labelling, images were collected independently.

For live-cell imaging, cells were grown as monolayers in 35 mm µ-dishes (Ibidi, Germany). Cells were pulsed with either HSA–AF488 or HSA^H464Q–AF488 in the presence of mCSF-1 for 5 min, washed three times with PBS, and Leibovitz's L15 medium (#21083027, Life Technologies) was added for imaging purpose. Image acquisition were performed using a Leica SP8 confocal system using a 63×/1.4 NA objective at 12 s intervals. Live-cell imaging results were analysed and prepared using ImageJ and TrackMate plugin.

Cells were prepared in RIPA buffer [50 mM Tris-HCl, pH 7.3, 150 mM NaCl, 0.1 mM EDTA, 1% (w/v) sodium deoxycholate, 1% (v/v) Triton X-100, 0.2% (w/v) NaF and 100 µM Na₃VO₄] supplemented with Complete mini protease inhibitors (Sigma Aldrich). Cell lysates were incubated on ice for 10 min and centrifuged for 10 min at 16,000 g. Protein concentrations were determined with a Bradford assay prior to SDS-PAGE. Proteins were resolved by SDS-PAGE (Life Technologies) and transferred onto Immobilon-P PVDF membrane (Millipore) at 30 V overnight at 4°C. The membrane was blocked by drying at 37°C. The membrane was incubated with primary antibodies diluted in 5% skim milk in PBS for 1 h and then washed three times for 10 min each in 0.1% (v/v) Tween 20 in PBS. HRP-conjugated secondary antibodies were then added to the membrane for 1 h and washed as above. Bound antibodies were detected through chemiluminescence and captured using the BioRad ChemiDOC imager (California, USA). Densitometry of the protein bands were measured using BioRad ImageLab program.

Cells were analysed at a medium flow rate in an LSRFortessa flow cytometer, equipped with 405 nm, 488 nm, 561 nm and 640 nm lasers (BD Biosciences). Approximately 10,000 events were collected, using a forward scatter threshold of 5000.

All analyses included samples from three or more independent experiments.

Data represent the mean±s.e.m. from three independent experiments, where for each experiment, an average value was determined from two to five images (each containing eight or more cells) for each time point and analysed by an unpaired two-tailed Student’s t-test. P<0.05(*) was considered significant, P<0.01 (**), highly significant and P<0.001 (***), very highly significant. An absence of a P value indicates that the differences were not significant.

Confocal microscopy was performed at the Biological Optical Microscopy Platform (BOMP) at the University of Melbourne. We gratefully acknowledge Fiona Houghton (Department of Biochemistry and Molecular BIology, Bio21 Institute, University of Melbourne) for expert technical advice, and Shirley Taylor for the generation of fluorescent conjugates.