Rationale.

Phage therapy is an investigational anti-infective treatment for refractory, multidrug-resistant (MDR), and/or biofilm-mediated infections. These situations include chronic and recurrent infections such as UTIs, rhinosinusitis, skin and soft-tissue infections, and MDR respiratory infections, as described below. Biofilm-associated infections include PJI, osteomyelitis involving hardware, cardiac device infection, and respiratory infections in the setting of cystic fibrosis (CF). In general, phage therapy should be limited to treatment of infections after intolerance or ineffectiveness of antibiotic therapy has been demonstrated. These scenarios generally occur in the setting of MDR infections and/or hardware-associated infections not amenable to source control. Phages with activity against Staphylococcus aureus and Pseudomonas aeruginosa are most readily available from academic as well as some commercial entities. Phages with activity against organisms such as Klebsiella pneumoniae, Enterococcus faecalis and faecium, Acinetobacter baumannii, Escherichia coli, Proteus mirabilis, Burkholderia species, and nontuberculous mycobacteria, among others, are less readily available but have been used to treat human infections. Phage therapy is currently limited to treatment of bacterial infections; however, data are emerging to indicate that nonbacterial organisms such as Aspergillus species can be treated in the future; this is not the subject of this work (1, 2). Members of the ARLG Phage Taskforce reviewed available data on experience with phage therapy for treatment of specific infection types. Table 1 summarizes published clinical experiences from case reports and case series on the use of phage therapy to treat human infections in recent years. Gleaning the potential benefit of phage therapy from the published literature is challenging for several reasons. First, most clinical publications do not include a control group. Second, there is heterogeneity in the route of administration, dose of phage, number of phages (i.e., monotherapy versus cocktail), and duration of phage therapy administered across studies. Third, phages are almost universally administered in addition to antibiotics, making an understanding of the specific role of phages in impacting clinical outcomes difficult to discern. Fourth, the use of phage susceptibility testing (PST) to ensure the phages administered are indeed active against the bacterial pathogen has been inconsistent and is nonstandardized (see Question 7). Finally, publication bias exists, with reports of unfavorable outcomes being less likely to be published than those with favorable outcomes. Below we summarize some of the most common infection types for which phage therapy has been used to date.

TABLE 1

Summary of recent published clinical reports and case series of phage therapy in humans^a

Article (reference); PMID	Organism	Demographics	Infectious syndrome	Route of phage administration	No. of phage administered	Highest dose of phage administered (PFU)	Frequency of phage administration	Duration of phage administration	Other anti-infectives administered simultaneously	Clinical outcomes	Survived initial infection	Adverse events	Follow-up period
Corbellino, 2020 (5); 31414123	Klebsiella pneumoniae	57-yr-old female	Urinary tract infection (from intestinal colonization)	Oral and intrarectal	1 phage	1×106	q12h	3wk	Yes	Remained free of infection with the colonization strain	Yes	No	11mo
Aslam et al., 2020 (7); 33005701	Staphylococcus aureus	61-yr-old male	Periprosthetic joint infection	i.v. and direct application	3 phages	3×109	i.v. q12h; intra-articular injection once	2wk	Yes	Clinical improvement at 2wk but recurrence on day 17 that necessitated additional antibiotics, phage therapy, and surgery	Yes	None reported	7mo
Aslam et al., 2020 (7); 33005701	Escherichia coli	56-yr-old male	Urinary tract infection	i.v.	4 phages	1×109	q12h	2wk	Yes	No clinical symptoms for 12wk but urine culture remained positive	Yes	None reported	12wk
Aslam et al., 2020 (7); 33005701	Pseudomonas aeruginosa	64-yr-old male	Aortic graft infection		3 phages	2.6×106	q12h	6wk	Yes	Clinical resolution	Yes	None reported	12wk
Kuipers et al., 2019 (8); 31611357	K. pneumoniae	58-yr old male	Urinary tract infection	Oral and direct application	Unknown	Unknown	q12–24h	16wk	Yes	Symptomatic relief	Yes	None reported	40wk
Bao et al., 2020 (9); 32212918	K. pneumoniae	63-yr-old female	Urinary tract infection	Direct application	5-6 phage cocktails	5×108	q24h	5days	Yes	Clinical cure	Yes	None reported	6mo
Khawaldeh et al., 2011 (10); 21737541	P. aeruginosa	67-yr-old female	Urinary tract infection	Direct application	6 phages	6×106	q12h	10days	Yes	Symptomatic relief; microbiological improvement with nearly 2-log reduction in P. aeruginosa urinary concentrations	Yes	None reported	12mo
Aslam et al., 2020 (7); 33005701	P. aeruginosa	82-yr-old male	Ventricular assist device infection	i.v. and direct application	4 phages	7.58×105	q8–12h	~10wk	Yes	Recurrent bacteremia within 1wk of completing phage therapy; retreated with phage with recurrent bacteremia 4wk into second episode while receiving phage	Yes	Fever, wheezing, shortness of breath after two infusions of 1011 PFU/mL concn; same phage well tolerated at 1010PFU/mL	5mo
Aslam et al., 2020 (7); 33005701	P. aeruginosa	60-yr-old male	Ventricular assist device infection	i.v.	3 phages	1.9×107	q8h	6wk	Yes	Developed bacteremia 1 wk after starting phage therapy; developed recurrent purulent drainage following end of phage therapy	Yes	None reported	6wk
Jikia et al., 2005 (17); 15663496	S. aureus	52-yr-old male	Skin and soft tissue infection	Direct application	≥5 phages	Not described	Once	Once	Yes	Clinical recovery within 7days	Yes	None reported	7days
Law et al., 2019 (20); 31102236	P. aeruginosa	26-yr-old female	Cystic fibrosis exacerbation	i.v.	4 phages	4×109	q6h	8wk	yes	Clinical improvement 7days into therapy; no CF exacerbations within 100days following end of phage therapy	Yes	None reported	100days
Gainey et al., 2020 (21); 32662948	Achromobacter species	10-yr-old female	Cystic fibrosis exacerbation in a cystic fibrosis patient	i.v.	1 phage	Not described	q24h	28days (14days without antibiotics and 14days with antibiotics)	Yes	FEV1 improved when administered with antibiotics; Achromobacter sp. could not be recovered up to 16wk after treatment	Yes	None reported	16wk
Hoyle et al., 2018 (22); 29777836	Achromobacter xylosoxidans	17-yr-old female	Cystic fibrosis exacerbation in a cystic fibrosis patient	Oral and nebulized	2 phages	3×108	q24h	20days	Yes	Dyspnea resolved and cough decreased; FEV1 increased from 54% to 84%	Yes	None reported	12mo
Dedrick et al., 2019 (23); 31068712	Mycobacterium abscessus	15-yr-old female	Pneumonia in a lung transplant recipient with disseminated lesions	i.v.	3 phages	1×109	q12h	~4mo	Yes	Clinical improvement, including resolution of infected skin nodules	Yes	Sweating and flushing for 2day	6mo
Aslam et al., 2019 (24); 31207123	P. aeruginosa	67-yr-old male	Pneumonia in a lung transplant recipient	i.v. and nebulized	3-5 phages	5×109	i.v. q2–6h and nebulized q6–12h	29days	Yes	New hospitalization for pneumonia on day 46 (with P. aeruginosa); recovered well from both episodes of pneumonia	Yes	None reported	29days
Aslam et al., 2019 (24); 31207123	Burkholderia dolosa	28-yr-old male	Pneumonia in a lung transplant recipient	i.v.	1 phage	3.5×107	q12–24h	12wk	Yes	Recovered	No	None reported	Died at 6mo from multiorgan system failure
Aslam et al., 2019 (24); 31207123	P. aeruginosa	52-yr-old female	Pneumonia in a lung transplant recipient	i.v.	4 phages	4×109	q12h	4wk	Yes	Recovered	Yes	None reported	60days
Dedrick et al., 2021 (25); 34239133	M. abscessus	81-yr-old male	Refractory pulmonary infection in the setting of bronchiectasis	i.v.	3 phages	1×109	q12h	6mo	Yes	Initial treatment response, followed by increased bacterial counts and ultimately treatment failure	Yes	None	NA
Wu et al., 2021 (26); 33703996	A. baumannii	62-yr-old male	Secondary pneumonia in COVID-19	Nebulized	1 phage, then 2 phages	1×109	2 administrations, 1day apart	2days	Yes	Decline in semiquantitative A. baumannii burden; clinical cure	Yes	Fever, IL-6, IL-8 cytokine storm; resolved after 1day	30days
Wu et al., 2021 (26); 33703996	A. baumannii	64-yr-old male	Secondary pneumonia in COVID-19	Nebulized	2 phages	1×109	2 administrations, 1hour apart	1day	Yes	Decline in semiquantitative A. baumannii burden; clinical cure	Yes	None reported	9days
Wu et al., 2021 (26); 33703996	A. baumannii	81-yr-old male	Secondary pneumonia in COVID-19	Nebulized	2 phages	1×109	2 administrations, 1hour apart	1day	Yes	Decline in semiquantitative A. baumannii burden; died day 10	No	None reported	10days
Wu et al., 2021 (26); 33703996	A. baumannii	78-yr-old male	Secondary pneumonia in COVID-19	Nebulized	2 phages	1×109	2 administrations, 1hour apart	1day	Yes	Decline in semiquantitative A. baumannii burden; died day 40 of K. pneumoniae infection	No	None reported	40days
Maddocks et al., 2019 (27); 31437402	P. aeruginosa	77-yr-old female	Pneumonia and empyema	Nebulized and i.v.	4 phages	1×109	q12h	7days	Yes	Clinical cure	Yes	None reported	6mo
Nir-Paz et al., 2019 (32); 30869755	A. baumannii, K. pneumoniae	42-yr-old male	Osteomyelitis	i.v.	2 phages	5×107	q8h	11days	Yes	Rapid tissue healing within a few days; wound healed by 2wk; prevented likely leg amputation	Yes	None reported	8mo
LaVergne et al., 2018 (33); 29687015	A. baumannii	77-yr-old male	Craniotomy site infection	i.v.	1 phage	2.14×107	q2h	8days	Yes	Direct improvement of craniotomy site	No	2 h after first dose became transiently hy potensive (no pressors)	Died at day 20 from withdrawal of care due to poor neurologic status
Cano et al., 2020 (34); 32699879	K. pneumoniae	62-yr-old male	Periprosthetic joint infection	i.v.	1 phage	6.3×1010	q24h	40days	Yes	Reduction in biofilm mass within 24h; improved mobility	Yes	None reported	8mo
Khatami et al., 2021 (35); 34369652	P. aeruginosa	7-yr-old female	Septic arthritis and osteomyelitis	i.v.	1 phage	1×1011	q12–q24	2wk	Yes	Clinical recovery	Yes	Fever, transient increase in heel pain	5mo
Onsea et al., 2019 (36); 31548497	P. aeruginosa, Staphylococcus epidermidis	Unknown	Osteomyelitis of femur	Direct application	2 phages	2×107	q8h	10days	Yes	Recurrence of infection 8mo later	Yes	None reported	16 mo
Onsea et al., 2019 (36); 31548497	Enterococcus faecalis	Unknown	Osteomyelitis of femur	Direct application	6 phages	Unknown	q8h	7days	Yes	Recovered	Yes	None reported	8mo
Onsea et al., 2019 (36); 31548497	P. aeruginosa, S. epidermidis	Unknown	Osteomyelitis of pelvis	Direct application	2 phages	2×107	q8h	7days	Yes	Recovered	Yes	Direct redness, pain	16mo
Onsea et al., 2019 (36); 31548497	Streptococcus agalactiae, S. aureus	Unknown	Osteomyelitis of femur	Direct application	2 phages	2×107	q8h	9days	Yes	Recovered	Yes	None reported	8mo
Ferry et al., 2020 (37); 32850878	S. aureus	49-yr-old male	Periprosthetic joint infection	Direct application	2 phages	2×1010	Once	1day	Yes	Recurrence of infection with different pathogens leading eventually to amputation	Yes	None reported	12mo
Ferry et al., 2018 (38); 30474047	S. aureus, P. aeruginosa (suspected but not proven)	80-yr-old female	Periprosthetic joint infection	Direct application	6 phages	6×1010	Once	1day	Yes	Favorable outcome	Yes	None reported	18mo
Ferry et al., 2018 (39); 30060002	P. aeruginosa	60-yr-old male	Osteomyelitis	Direct application	4 phages	(1.2–9.7)×108	Every 3 days for 4h at a time	12days	Yes	Gross appearance of wound improved by day 14	No	None reported	Died at day 45 due to spine metastases
Tkhilaishvili et al., 2019 (40); 31527029	P. aeruginosa	80-yr-old female	Periprosthetic joint infection	Direct application	1 phage	1×108	q8h	5days	Yes	No P. aeruginosa isolated in joint fluid samples in first 5days and 4wk after phage instillation and up to 10mo later	Yes	None reported	10mo
Ferry et al., 2021 (41); 34026768	P. aeruginosa	88-yr-old male	Periprosthetic joint infection	Direct application	2 phages	3×1010	Once	Once	Yes	Resolution	Yes	None reported	1yr
Ferry et al., 2020 (42); 33304911	S. aureus	80-yr-old male	Periprosthetic joint infection	Direct application	3 phages	1×109	Once	Once	Yes	Resolution	Yes	None reported	2yr
Ferry et al., 2020 (42); 33304911	S. aureus (suspected but not proven)	84-yr-old male	Periprosthetic joint infection	Direct application	3 phages	1×109	Once	Once	Yes	Resolution	Yes	None reported	7mo
Doub et al., 2021 (43); 33800146	S. epidermidis	79-yr-old female	Periprosthetic joint infection	Direct application	1 phage	2×1010	Once	Once	Yes	No recurrence on chronic suppresion; resolution of chronic anemia	Yes	Trans ient elevation in liver enzymes	5mo
Ferry et al., 2020 (43); 33304911	S. aureus	83-yr-old female	Periprosthetic joint infection	Direct application	3 phages	1×109	Once	Once	Yes	Pain-free and functional joint but with intermittent drainage from fistula	Yes	None reported	11mo
Ramirez-Sanchez et al., 2021 (44); 34205687	S. aureus	61-yr-old female	Periprosthetic joint infection	i.v. and direct application	1 phage	3×1010	q12h	6wk	Yes	Clinical cure	Yes	None reported	20mo
Doub et al., 2020 (45); 32397354	S. aureus	72-yr-old male	Periprosthetic joint infection	i.v. and direct applplication	1 phage	5×109	q24h	3days	Yes	Resolution	Yes	Transient elevation in liver enzymes	2mo
Exarchos et al., 2020 (46); 31740948	S. aureus, Cutibacterium acnes	41-yr-old man	Cardiovascular implantable electronic device and aortic graft infection	Direct instillation	2 phages	1×106	q8h	14days	Yes	Clinical cur e	Yes	None reported	12mo
Aslam et al., 2019 (47); 30661974	S. aureus	65-yr-old male	Ventricular assist device infection	i.v.	3 phages	3×109 i.v.	q12h	28days	Yes	Clinical improvement within 1wk; continued to be well appearing 7mo later	Y es	None reported	7mo
Mulzer et al., 2020 (48); 31651936	S. aureus	67-yr-old male	Ventricular assist device infection	Direct application	2 phages	2×107	q8h	10days	Yes	Clinical cure	Yes	Mild nausea	9mo
Tkhilaishvili et al., 2021 (49); 34058260	P. aeruginosa	53-yr-old male	Ventricular assist device infection	i.v. and direct applplication	3 phages	5×109	i.v. 7-h infusion direct q12h	5days	Yes	Clinical cure; patient expired 4mo later due to noninfectious cause	Yes	Mild nausea	4mo
Rubalskii et al., 2020 (50); 32380707	S. aureus	45-yr-old male	Infusion pump infection	Direct application	1 phage	4×1010	Once	Once	Yes	Clinical cure	Yes	None reported	>2yr
Rubalskii et al., 2020 (50); 32380707	Escherichia coli	66-yr-old female	Sternal infection after cardiac surgery	Direct application	2 phages	4×1010	Once	Once	Yes	Eradication of E. coli	Yes	None reported	2yr
Rubalskii et al., 2020 (50); 32380707	P. aeruginosa	13-yr-old male	Sternal infection after lung transplant	Direct application	2 phages	4×1010	Once	Once	Yes	Resolution	Yes	None reported	2yr
Rubalskii et al., 2020 (50); 32380707	S. aureus	51-yr-old male	Ventricular assist device infection	Direct application and oral	4 phages	1×109	q12–24h	12days	Yes	Microbiologic reduction of S. aureus	No	None reported	6wk
Rubalskii et al., 2020 (50); 32380707	S. aureus	59-yr-old male	Aortic graft infection	Direct application	1 phage	1×109	q12h	2days	Yes	Clinical cure	Yes	None reported	>2yr
Rubalskii et al., 2020 (50); 32380707	S. aureus	62-yr-old male	Ventricular assist device infection	Direct application	1 phage	1×109	q12h	7days	Yes	Clinical cure; died 20mo after heart transplant due to transplant failure	Yes	None reported	20mo
Rubalskii et al., 2020 (50); 32380707	K. pneumoniae	40-yr-old male	Pneumonia in settning of heart transplant	Nebulized and oral	2 phages	1×108	q12–q24h	4days	Yes	Clinicalcure	Yes	None reported	>3yr
Rubalskii et al., 2020 (50); 32380707	S. aureus, Enterococcus faecium, P. aeruginosa	52-yr-old male	Aortic graft infection	Direct application and oral	4 phages	1×108	One dose orally, direct instillation, and intraoperatively	1day	Yes	Staphylococcus aureus, Enterococcus faecium, Pseudomonas aeruginosa not detected but died at 2mo from a new bacterial infection	Yes	None reported	4days
Chan et al., 2018 (51); 29588855	P. aeruginosa	76-yr-old male	Aortic graft infection	Direct application	1 phage	1×107	Once	Once	Yes	Recovered	Yes	None reported	18mo
Gilbey et al., 2019 (52); 31281964	S. aureus	65-yr-old male	Periprosthetic valve endocarditis	i.v.	3 phages	3×109	q12h	14days	Yes	Recovered and discharged home but on day 98 had progressive heart failure and a new potential vegetation with sterile blood cultures; declined surgery and died on day 103	Yes	None reported	103days
Schooley et al., 2017 (54); 28807909	A. baumannii	68-yr-old male	Necrotizing pancreatitis with infected pancreatic pseudocysts	i.v. and direct application	9 phages (not all administered at the same time)	5×109	q6–8h	~12wk	Yes	Recovered	Yes	None reported	>2yr
Duplessis et al., 2018 (55); 28992111	P. aeruginosa	2-yr-old male	Endovascular infection and bacteremia	i.v.	2 phages	3.5×105	q6	6days	Yes	Sterilization of blood cultures; patient expired due to cardiac and septic shock attributed to undrained fluid collections	No	Decompensation attributed to heart failure but endotoxin release could not be excluded	None
Jennes et al., 2017 (57); 28583189	P. aeruginosa	61-yr-old male	Infected pressure sores	i.v. and direct delivery (50mL phage solution irrigation)	2 phages	2×106	i.v. q6h; direct delivery q8h	10days	Yes	Pressure sores remained infected with intermittent episodes of sepsis	No	None reported	Died at 4mo from Klebsiella pneumoniae sepsis
Fadlallah et al., 2015 (58); 26862360	S. aureus	65-yr-old female	Bacterial keratitis	Direct application	Not described	Not described	q12h	4wk	Yes	Clinical resolution and negative ocular cultures	Yes	None reported	6mo
Johri et al., 2021 (62); 34177601	P.	33-yr-old male	Prostatitis	Oral liquid, rectal suppositories, and urethral instillations	≥3 phages cocktails	Not described	Daily oral; BID suppositories; daily urethral	14days oral; 10days rectal; 10 d ays urethral followed by 2mo of oral and rectal	No	Clinical and microbiological resolution	Yes	None reported	5mo
Fish et al., 2018 (63); 30279396	S. aureus	63-yr-old female	Osteomyelitis	Direct application	1 phage	1×109	Weekly	7wk	Yes	Partial improvement	Yes	None reported	3yr
Marza et al., 2006 (64); 16781080	P. aeruginosa	27-yr-old male	Skin graft infection	Direct application	1 phage	1×103	Once	Once	Yes	Clinical improvement and negative cultures	Yes	None reported	3days

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^aq12h, every 12 h; NA, not applicable; IL-6, interleukin-6.

Recurrent UTIs.

There are anecdotal, uncontrolled reports of successful use of phage therapy (in combination with antibiotics) for treatment of patients with recurrent UTIs. Gut, vaginal, and urinary microbiomes are important reservoirs of uropathogens and likely contribute to the pathophysiology of recurrent UTIs (3, 4). In a patient with recurrent UTI secondary to MDR-K. pneumoniae, oral and intrarectally administered phage resulted in microbiologic clearance in urine and stool (5). In addition to potentially decreasing the gut reservoir of uropathogens, phages may decrease uropathogen colonization of urinary catheters. In vitro data suggest that phage-treated catheters can be protected from colonization with common uropathogens (6). Other case reports showing successful treatment of UTI using phages include a liver transplant recipient with recurrent extended-spectrum beta-lactamase (ESBL)-producing E. coli UTIs (7), a kidney transplant recipient with an ESBL-producing K. pneumoniae UTI (8), an extensively drug-resistant (XDR) K. pneumoniae UTI (9), and a refractory P. aeruginosa UTI in the setting of ureteral stents (10). A single randomized controlled trial investigating phage therapy in men presenting for transurethral resection of the prostate with complicated or recurrent UTI yielded unfavorable results (11). In this trial, 28 patients received intravesicular phage, 32 received placebo mechanical irrigations, and 37 received systemic antibiotics, with no differences in urine culture sterilization across the three groups. Phage therapy was not inferior to standard-of-care antibiotic treatment, and it also was not superior to placebo bladder irrigation. Absolute success rates were lowest in the phage (18% success) compared to the placebo (28% success) and antibiotic (35% success) groups. Low concentrations of phages (10⁴ to 10⁵ PFU per dose) and insufficient coverage of bacterial pathogens due to use of a fixed cocktail and/or poor phage survival in urine (influenced by pH and temperature) may have impacted this study.

Chronic rhinosinusitis and otitis media.

Chronic and recurrent bacterial rhinosinusitis can be difficult to cure with conventional antibiotic approaches secondary to biofilm formation and pathogens with difficult-to-treat resistance patterns (12). In a pilot uncontrolled clinical trial of nine patients with chronic rhinosinusitis, intranasal phages were administered for up to 14days. Treatment was well tolerated, and infection was eradicated in two patients (12). A phase I/II double-blinded, placebo-controlled trial was conducted in 2009 randomizing 24 patients with drug-resistant chronic refractory P. aeruginosa otitis media into phage or placebo arms; three patients in the phage arm had undetectable P. aeruginosa compared with none in the placebo group (13). A follow-up phase III study has not been conducted.

Skin and soft-tissue infections.

Like rhinosinusitis, skin infections offer the possibility for topical phage administration (14, 15). However, in a clinical trial including 13 patients randomized to phage therapy alone and 14 randomized to standard of care alone, phage decreased the bacterial bioburden but at a slower rate than standard of care in P. aeruginosa-infected burn wounds. Low phage concentrations, lack of baseline PST, and poor adherence of phage to the dressings used may have contributed to the negative trial results (16). Radiation burn wounds infected with S. aureus (a highly specific infection type) were successfully treated with topical phage (Table 1) (17).

Respiratory infections.

Recurrent infections with MDR Gram-negative bacilli are common in patients with CF as well as lung transplant recipients. There are case reports of phage therapy in CF patients with respiratory infections caused by MDR Gram-negative bacteria (18,–22) (Table 1). In one such case, a patient with CF and worsening acute-on-chronic respiratory failure due to MDR P. aeruginosa despite antibiotic therapy improved with concomitant addition of four intravenous (i.v.) phages for 8 weeks, enabling the patient to undergo successful bilateral lung transplantation (20). Hoyle et al. published the experience of a teenager with CF and an MDR Achromobacter spp. pulmonary infection who received both nebulized phages once daily as well as oral phage twice daily periodically over a year, with improved lung function (22). Several other cases of apparent successful use of prolonged courses of i.v. and inhaled phage to treat respiratory infections in lung transplant patients have been reported (23,–25) (Table 1). For example, a 15-year-old female with CF who had undergone bilateral lung transplantation and developed disseminated M. abscessus infection, including skin and soft-tissue and respiratory infection, was successfully treated using a three-phage cocktail, including an engineered phage, in addition to traditional anti-infective therapy (23). However, clinical failure was observed in an immunocompetent 81-year-old patient with refractory M. abscessus pulmonary disease in the setting of bronchiectasis, associated with development of a strong neutralizing antibody response (see Question 9) (25). A series of four patients with coronavirus disease 2019 (COVID-19) and secondary infection with A. baumannii were treated with nebulized phages, with clinical cure in two cases (26) (Table 1). Another non-CF case involved a patient with refractory P. aeruginosa pneumonia and empyema treated with i.v. and nebulized phages for 7days, with clinical cure (27).

Bone and joint infections.

Bacteria in biofilms possess a variety of mechanisms for immune and antibiotic evasion, making biofilm-related infections both difficult to detect and eradicate. Phages are a potential approach for treating biofilm-mediated musculoskeletal infections, especially when hardware cannot be removed. In vitro data indicate that some phages are active against biofilms of different ages (28) and with different extracellular matrix types (29).

Prior to the 21st century, there were several case series, primarily from Poland and the Republic of Georgia, describing use of phages for treatment of musculoskeletal infections, generally in conjunction with antibiotics, with clinical success described in upwards of 90% of cases (30, 31). Several clinical phage treatment experiences with musculoskeletal infections from diverse parts of the world have been published in recent years (Table 1). Specific musculoskeletal infections for which phage therapy has been used include PJI, spinal hardware infection, trauma-related injury associated with infected hardware, and craniectomy-related infection. Phage therapy has been administered both i.v. (7, 32,–35) and locally to treat musculoskeletal infections (36,–40). To highlight one case series, Onsea et al. describe their experience using phage therapy to treat four orthopedic infections, using a standardized treatment approach to locally administer phage (36). Phages were administered intra- and postoperatively three times daily for a maximum of 10days. All patients also received antibiotics; their clinical status was monitored daily during phage therapy. There were no recurrences of infection over 8 to 16months of follow-up (36).

PJI.

Several case reports employed local instillations of phages in patients with chronic S. aureus, Staphylococcus epidermidis, or P. aeruginosa PJI, with phages administered in conjunction with debridement, antibiotics, and implant retention (DAIR). This approach has yielded successful outcomes (Table 1) in patients who were not candidates for resection arthroplasty, allowing implant salvage while controlling or curing the infection (37,–39, 41,–43). Another case reported clinical cure of chronic K. pneumoniae PJI with an i.v. phage without surgical debridement (34). Other reports describe successful treatment of chronic S. aureus PJI with resection arthroplasty (44, 45).

Cardiac device-associated infection.

Cardiac infections for which phage therapy have been administered include infections associated with cardiovascular implantable electronic devices (CIEDs) (46), ventricular assist devices (VADs) (7, 47,–50), vascular grafts (50, 51), and prosthetic valves (52). Several reports of CIED infections treated with phages have been described, generally due to S. aureus or P. aeruginosa (Table 1). A series of eight cases reported by Rubalskii et al. included infected vascular grafts, infected VADs, and sternotomy infections postcardiac surgery, caused by S. aureus, K. pneumoniae, E. coli, and P. aeruginosa; clinical resolution was achieved in seven of eight cases (50). In other reports, two cases of S. aureus and one case of P. aeruginosa VAD infection were successfully resolved with phage therapy in conjunction with antibiotics (47,–49); one patient with S. aureus infection was treated with i.v. phages for 4 weeks in addition to systemic antibiotics and the other with local instillation of phage through an indwelling drain for 10days, alongside systemic antibiotics, following debridement. The patient infected with P. aeruginosa was given a 7-h i.v. infusion of phages, followed by local application during surgical debridement, and then local instillation through an indwelling drain every 12h for 5days. Another patient with extensive local CIED infection caused by S. aureus associated with a bypass graft infection was successfully treated with debridement followed by local phage instillation for 2 weeks (46). However, two patients with P. aeruginosa VAD infections treated with i.v. phages for six to eight weeks, along with i.v. antibiotics, experienced infection recurrence (7). Gilbey et al. reported treating a patient with S. aureus aortic valve endocarditis with i.v. phages for 14days in addition to antibiotics; he recovered and was discharged but returned 3months later with progressive heart failure, potential vegetations, and negative blood cultures (52).

Sepsis.

As with all infections, the role of phage therapy in acute sepsis is unclear. In 2003, Weber-Dabrowska et al. reported observations in 94 septic patients failing antibiotic therapy with a mixture of monomicrobial and polymicrobial infections from a variety of syndromes, including UTI, skin and soft-tissue infection, respiratory infection, and intra-abdominal infection (53). Phage therapy was generally administered orally three times per day for a median of 29days and was given with antibiotics in 71 subjects. Recovery was achieved in approximately 85% of cases; there was no statistically significant difference between the antibiotics plus phage and phage monotherapy groups (53). In a separate study, a 68-year-old diabetic patient received phage therapy to treat necrotizing pancreatitis complicated by an MDR A. baumannii-infected pancreatic pseudocyst. The patient, who was comatose and septic, was successfully treated with phage therapy administered both i.v. and via percutaneous catheters directly into the infected cavities, showing significant improvement within 48h of phage administration (54). A 2-year-old patient with DiGeorge syndrome (Table 1) who had a mycotic aneurysm due to P. aeruginosa was given i.v. phage therapy with sterilization of blood cultures but ultimately succumbed to sepsis, attributed to undrained fluid collections and lack of surgical source control (55).

Rationale.

Recent clinical data on phage therapy has been generated primarily in compassionate use settings in conjunction with antibiotic therapy (7, 9, 20,–27, 32,–35, 37,–39, 41,–47, 50, 51, 54, 56,–58). Many cases were associated with apparent successful response of MDR and/or biofilm-associated infections that were not resolving with antibiotics alone, suggesting that in those cases, there may have been an additive or synergistic effect of the phage-antibiotic combination (9). An additional benefit of using phages in combination with antibiotics is the potential to reinstate susceptibility of the targeted bacteria to antibiotics by manipulating bacterium-phage coevolutionary strategies. In one study, the investigators collected bacterial isolates before and after phage treatment and observed changes in antibiotic susceptibility patterns and a reduction in pathogen fitness after treatment with phages (54). Chan et al. used a phage known to effect an evolutionary trade-off between phage resistance and antibiotic susceptibility in vitro (59) to successfully treat a patient with a P. aeruginosa-infected aortic graft (51). These clinical findings are supported by in vitro and animal model data demonstrating phage-antibiotic synergy in some circumstances (see Question 8).

Rationale.

Phages do not have reliable activity against all strains of any bacterial species, underscoring the potential importance of PST to identify phages active against the infecting pathogen before administration. Once standardized PST methodologies become available, members of the ARLG Phage Taskforce suggest that attempts be made to perform PST before phage administration where possible. Furthermore, because of the potential for resistance to develop to phage during treatment (26, 54), confirmation of the continued activity of phages against the bacterial pathogen is likely to be helpful to determine whether clinical failures may be due to emergence of phage resistance or alternative reasons, such as a need for source control. Evidence suggests that infections caused by pathogens that develop resistance to a phage cocktail are not effectively treated unless a new phage or phage cocktail that shows activity against the resistant isolate is administered (26, 54).

PST may involve testing of a panel of phages (sometimes referred to as generating a phagogram [7]) to select one or more phage for therapeutic use. Current approaches to determining phage susceptibility are described in Question 7. There is potential dynamic tension with the time testing may take to perform, especially as there is no available method that can be easily incorporated into clinical microbiology laboratory workflows and because testing may need to occur offsite. Phage cocktails or phages with broad host ranges may allow empirical phage therapy in some emergency situations; however, confirming phage activity is still considered ideal before initiation of therapy, once suitable PST methods become available.

Rationale.

Members of the ARLG Phage Taskforce reviewed common laboratory methods used for PST (sometimes referred to as host range testing), which have been applied to identify lytic phages potentially suitable for phage therapy. Two common approaches are double-layer agar and liquid testing methods.

With the double-layer agar method, a bacterium-phage mixture in melted low-concentration agar is spread over a solid agar nutrient medium in a petri dish (81). The semisolid state of the bacterium-phage layer restricts movement of bacteria and phages. The plate is incubated at a defined temperature for a defined period, dependent on the bacterial species tested (e.g., 18 to 24h), although such criteria have not been standardized. Bacteria multiply, producing a confluent lawn of bacteria in the top agar layer. Lytic phages can infect the bacteria, replicate, cause cell lysis, and produce progeny, which infect neighboring bacterial cells. The cycle of infection and lysis continues, killing bacteria in localized areas and ultimately creating plaques, with each plaque typically being the result of infective proliferation of one lytic phage (82). While larger clear plaques may represent greater effects than smaller plaques, plaque size may vary by phage and/or host. Turbid plaques may result from slight growth or formation of lysogens (Question 10). Limitations to this method include poor reproducibility, protracted turnaround time (83, 84), and inability to assess individual members of a phage cocktail (if tested together). Further, microbiological detection of phage activity only reflects the number of phages able to infect the host under select time constraints and to generate plaques visible to the naked eye, potentially missing phage subpopulations with clinically relevant biological or immunologic properties, such as phage immunoreactivity (85). Ion and metal content, agar concentration, the presence of organic compounds or detergents, specific antibodies, complement system elements, other phages, bacterial host age and growth phase, incubation temperature, and storage vessel may affect a phage’s infection capability (84,–87). The spot test is a modification of the double-layer agar method that involves spotting a phage suspension onto a solidified agar base over which bacteria in melted low-concentration agar has been placed (88). After overnight incubation, susceptibility to phage is determined by observing plaques at sites of spotting. A limitation of this approach is that plaques on the spot test can result from “lysis from without” due to phage lysins in phage lysate material and therefore may not necessarily reflect phage propagation (or “lysis from within”) (89). Although these methods require minimal instrumentation and equipment, they can be time and material intensive.

Another approach to PST involves liquid testing, with phage added to a bacterial inoculum, ideally at a defined ratio (known as the multiplicity of infection [MOI]). Liquid-based testing captures longitudinal, semiquantitative data addressing phage-mediated changes to bacteria (e.g., bacterial metabolism or bacterial abundance assessed by optical density). Susceptibility is determined by comparison to behavior of the bacterium without phage. This approach may be amenable to standardized, automated, scalable testing of phage activity and theoretically may be used to simultaneously test antibiotics and even to assess synergism or antagonism of combination therapies. Liquid-based assays furnish time-based measurements; theoretically, the longer the phage affects a bacterium, the greater the phage effect and/or the lower the chance for emergence of phage resistance may be. If this is the case, the ideal length of time needed to carry out such assays needs to be defined. That said, these theoretical possibilities have yet to be shown to be clinically relevant.

Rationale.

As discussed in Question 2, most recent clinical data on phage therapy have been generated when used in conjunction with antibiotics, making it difficult to draw conclusions as to the relative contributions of phages and antibiotics to outcomes. In vitro studies indicate that some phages lower some antibiotics’ MICs (96, 97), with phage-antibiotic synergy demonstrated under certain circumstances (98,–100). Thus, methods for ascertaining combination activity may inform selection of specific phage-antibiotic combinations.

A systematic assessment of activity of phages combined with antibiotics against E. coli revealed diverse responses, ranging from additive to synergistic to antagonistic effects, dependent on antibiotic class and resistance profile of the bacterial strain (9, 101). Some studies examining phage-antibiotic interactions in the presence of host factors such as urine and bile salts demonstrate synergy (102), whereas others do not (101, 103). Animal studies have demonstrated additive or even synergistic effects of certain phage-antibiotic combinations (104,–107). Phage-adjuvanted antibiotic effects were observed in an E. faecalis sepsis model, suggesting that, under some circumstances, lower doses of antibiotics are acceptable when combined with phages (104). In the case of antagonism, phage sequencing may be useful to compare the similarity of ribosomal genes or DNA polymerase to those of the bacterial host; if they are similar, antibiotics that inhibit these may impact the phage life cycle.

There is evidence that phage-antibiotic combinations can reduce the likelihood of emergence of antibiotic- and phage resistant-bacteria and decrease bacterial fitness (54). In a study using over 400 A. baumannii isolates, there was a correlation between antibiotic resistance and phage susceptibility, suggesting evolutionary trade-offs (108). Phage-plus-antibiotic combinations reduced phage and/or antibiotic resistance of S. aureus, E. coli, P. aeruginosa, and Enterococcus species in vitro (101, 109,–114). Phage specificity is related to phage-receptor interactions (115), with internal bacterial cellular functions (e.g., gene expression and DNA replication) also playing a role (115). Some phages use virulence factors (116) or antibiotic resistance proteins (59) as receptors, such that bacteria that develop resistance to the phages via receptor mutation may be less virulent or more antibiotic susceptible, respectively.

Phage-antibiotic synergy has also been observed in biofilms. When tested against monospecies or mixed biofilms, phages combined with antibiotics reduced bacterial densities of P. aeruginosa, S. aureus, and enterococcal biofilms in vitro (117,–121), with enhanced activity if treatment with phages preceded antibiotics (98, 122). Similar results were observed against P. aeruginosa from CF airways and wounds (123). In an animal model of orthopedic implant-related infection, S. aureus but not P. aeruginosa biofilm thickness was reduced following phage-antibiotic combinations more so than either alone, suggesting that bacterial species, alongside infection type or site, impacts effects of combination activity (124).

Emerging methods might consider consolidating phage and antibiotic susceptibility testing methods in a way that could allow for combination testing, for example, using a checkerboard assay in which various antibiotics are added to wells of a liquid testing plate (see Question 7); this may allow exposure of targeted bacteria to variable concentrations of antimicrobials and single or multiple phages (16, 125, 126). Gu Liu et al. offer a method for phage-antibiotic testing using 96-well plates that could be developed for use in clinical microbiology laboratories (101) to assess phage-antibiotic antagonism or synergy. Results of such testing need to be correlated with successful versus unsuccessful outcomes in clinical trials. Modified disk diffusion methods and time-kill analyses have been used to assess potential phage-antibiotic synergy (127).

Rationale.

Although bacteria are the designated hosts of phages, a complex interplay between the human (superhost) immune system, phages, and bacteria likely impacts phage activity and potentially larger aspects of human health and disease (128, 129). Few reports in the English literature have explored the nature of the immune response to phage therapy, although there is some evidence of synergy between the innate immune system and phages during therapy, with phagocytosis possessing a major role. Early work showed phages to be cleared by the reticuloendothelial system, with deposition in the spleen (130). A recent study showed lower phage concentrations in spleens of mice with extant LPS-induced systemic inflammatory responses and more effective phage inactivation by splenocytes from these compared to control mice (87). Roach et al. reported that neutrophils and innate immune cell signaling impact effective phage treatment of acute pneumonia in mice (131). In a recent report on the treatment of a 7-year-old girl with P. aeruginosa septic arthritis and osteomyelitis (see Question 1), there was upregulation of genes associated with innate and adaptive immunity in response to i.v. phage therapy (35).

Phages have been observed to reduce markers of inflammation, such as C-reactive protein, with treatment (132). Results of a recent study of severe S. aureus infection, including infective endocarditis and septic shock in humans, suggest that phage therapy induces an anti-inflammatory response; inflammatory markers decreased in 11 of 13 patients, and transcriptome analysis indicated changes in regulation of genes involved in the innate immune response in the blood of infective endocarditis patients following phage administration (56). However, a separate study of cytokine expression following human monocyte-derived dendritic cell treatment with S. aureus phage K found that phages had little impact on pro- and anti-inflammatory cytokine or CD80/CD86 and major histocompatibility complex class I/II protein expression (133). Because phage preparations may contain remnants of bacteria used for phage propagation, it may be difficult to distinguish immune responses elicited by phages from those elicited by bacterial components in phage preparations. A reasonable approach is to consider monitoring C-reactive protein with prolonged phage therapy, especially systemic administration.

Antibody responses to phages have been reported to develop over the course of phage therapy. Factors that contribute to levels and classes of antibodies generated include the route, dose, and frequency of administration. Studies in mice and humans have found weak antibody responses to be induced after oral administration (134,–136). Majewska et al. found that although phage-specific IgG antibodies were detected in blood early during prolonged oral phage treatment, only secreted IgA in feces, detected in late treatment, coincided with decreased phage levels (136). Arguably, antibodies may not be highly relevant to topical phage applications. In mice treated intraperitoneally (i.p.) with phages, antibody responses appeared to have an impact on phage levels, with a decrease in circulating phages in mice immunized with phages before subsequent treatment (87, 137). However, despite decreased phage levels, treatment was nonetheless effective in reducing bacterial burden and wound size in a murine model, albeit not to the extent of unimmunized treated mice (137). Similarly, the presence of phage-specific antibodies may not impede effective therapy in humans, as reported in 20 patients who received an antistaphylococcal phage cocktail (135). On the other hand, Dedrick et al. recently reported on a patient with bronchiectasis and M. abscessus pulmonary infection (see Question 1) who initially responded to an i.v. 3-phage cocktail with decreased M. abscessus counts in sputum but, with continued treatment, developed a neutralizing antibody response that corresponded to increased disease burden and treatment failure (25). The ARLG Phage Taskforce suggests considering measurement of neutralizing antibodies with prolonged courses of phage therapy but recognizes that no standardized assays exist to measure neutralizing antibodies.

Rationale.

As with any medical intervention, the first consideration is safety. Primary safety concerns include phage predilection to lysogeny and formulation sterility and mitigation of secreted bacterial products.

The history of the phage and bacterial propagation host should be well-documented. Ideally, annotated whole-genome sequences of the phage and host bacterial strain used for its propagation should be used to interrogate for known therapeutically deleterious elements (e.g., antibiotic resistance elements, toxin genes, prophages, integrase genes, regulators of integrase genes, and integrase-like genomic elements) in their genomes. Sequenced bacterial propagation hosts that do not produce toxins or harbor antibiotic resistance genes are preferred. Absent a trusted bacterial propagation host, the patient isolate itself may be considered a phage propagation host. In such cases, the isolate could be sequenced to show that it does not harbor toxin genes or antibiotic resistance genes, although given that phage therapy is often targeted at drug-resistant bacteria, this may not be possible. If there is no time to sequence the patient isolate’s genome, testing for specific toxins characteristic of the species (e.g., Shiga toxin for E. coli, toxic shock syndrome toxin 1 for S. aureus) may be considered, but it should be noted that bacterial species may encode toxins traditionally harbored in other species because of genetic exchange.

Phages may mediate transfer of genes to bacteria by lysogeny (or generalized transduction). Lysogeny (also known as lysogenic conversion) involves integration of a temperate phage genome into a bacterium’s genome. Phages that undergo lysogenic conversion are not ideal for therapeutic use, as they can render bacteria more virulent if the phage harbors deleterious genes. The ability of phages to undergo lysogeny may be assessed phenotypically and genotypically, with neither method being perfect. Genotypic methods involve sequencing phage DNA and analyzing sequence data for genomic elements involved in lysogeny (i.e., integrases, enzymes that mediate incorporation of phage DNA into bacterial DNA; integrase-like genomic elements; and regulators of lytic genes, such as repressor genes that help maintain prophages in a quiescent state). Phage genomes can also be examined for homology to known temperate phages, which suggests a lysogenic lifestyle. For example, a bioinformatics tool for computational evaluation of phage lifestyle such as the Phage Classification Tool Set (PHACTS) can be used (139). Integrase genes are diverse, so relying on sequence similarity to published genomes may be insufficient to accurately predict phage lifestyle. The annotation tool PHASTER, for example, detected only half of 147 integrase genes in prophages from 49 Salmonella enterica isolates (140). In addition, integrase genes are not necessary for a lysogenic lifestyle, with some phages replicating as plasmids in the lysogen. Further, mutations in single amino acid residues may render integrases inactive (141); thus, functional capacity of an integrase cannot be definitively known unless it is identical to an experimentally confirmed annotation. PHACTS does not universally provide clear classifications, and some differences are explained by specialized host-phage interactions that govern lifestyle by way of interference of integrases or other repressor genes (142). Philipson et al. recently described a multiplexed bioinformatics workflow to produce a fully assembled phage genome and to demonstrate that phages appear to be lytic and to not encode therapeutically deleterious genes (143). They propose that generated phage genomes be considered “finished” when they contain a single consensus sequence representing 100% of the genome with all open reading frames identified and a lack of population diversity, indicating purity of the sequence and verified via deep sequence coverage. PhageTerm is an example of a tool for determining genomic termini and phage packaging strategy (144). Sequences are evaluated for toxins and resistance genes, host, and laboratory contamination.

For phenotypic testing, phage plaques may be visualized to assess turbidity, which suggests that lysogeny may have occurred. This method is neither perfectly sensitive nor specific; even if plaques are clear, lysogeny may have happened but not be noticeable by eye. A sterile pipette tip touched to the plaque may be streaked on a plate of appropriate medium; bacteria that grow may be lysogens. These colonies can be retested for resistance to the phage; if resistant, they can be confirmed as lysogens if phages are detected after spontaneous excision or following induction with bacterial stressors or mutagens (e.g., mitomycin C, UV radiation, carbadox, peroxide, temperature stress). Caution is needed, however, as spontaneous phage resistance coupled with imprecise technique carrying over phage particles from plaques could appear to be a lysogen by this approach. Alternately, the genome sequence of a putative lysogen could be examined for the presence of a prophage absent from the parent. If the ability of a particular phage being evaluated to undergo lysogeny varies with the host bacterium assessed, definitive prediction of absence of lysogeny may be impossible.

Generalized transduction is another mechanism by which phages can mediate genetic transfer as segments of host DNA become incorporated into phage capsids by chance during the assembly phase of replication. Generalized transduction is an undesirable potential outcome of phage therapy, although it is impossible to completely mitigate. When recombinant phages encounter subsequent hosts, DNA may be incorporated into the bacterial genome through homologous recombination, resulting in transfer of DNA from one bacterium to another. Because of the stochasticity of phage assembly, most chromosomal sequences are transduced with roughly equal frequency.

Regarding phage formulation, preparations should be labeled with information on phage identity, purity, strength, stock expiration date, and storage conditions (115). An issue with phage stability is the occurrence of mutations that can impair viral fitness in phage stocks stored for long periods or accumulated during manufacturing and phage production. Development of manufacturing processes that minimize mutation of phage genomes is needed. Phage preparations should also be free of viable bacteria and fungi and contain minimal amounts of bacterial secreted products/debris (e.g., spores, endotoxin, exotoxins). Due to their classification as biological therapeutics, phage products used in the United States need to be manufactured under current good manufacturing practice (GMP) and adhere to USP requirements based on the type of application before administration. FDA guidance for GMP for phase I investigational products is available (145). Endotoxin levels in phage products need to be below acceptable limits set by the FDA and vary based on the route of administration. Methods to quantify endotoxin have been described (146); endotoxin requirements are <0.5 endotoxin units (EU)/ml for subcutaneous injections, <5 EU/kg of body weight/h for i.v. injections, and <0.2 EU/kg for intrathecal injections (147, 148). In industry and molecular biology applications, phage purification has often been carried out using polyethylene glycol precipitation followed by serial chloroform extractions, ultracentrifugation on a cesium chloride (CsCl) gradient, and subsequent dialysis to remove CsCl. Chloroform concentrations are regulated in medical products; therefore, the use of chloroform necessitates analysis of residual solvent concentration. CsCl is a theoretical safety risk (149), although residual CsCl concentrations in phage products would be unlikely to be significant (150). Other methods have been described to deplete endotoxin from phage preparations, such as polyethylene glycol, ultrafiltration, ultracentrifugation, gel filtration, anion-exchange chromatography, octanol extraction, deoxycholate extraction, endotoxin removal columns, and combinations thereof (151,–156); not all work equally well for all bacterial species. Phage purification studies to date have mostly focused on phage yield and endotoxin removal, with little focus on removal of other bacterial toxins during phage purification. Gram-positive bacteria do not have endotoxin, but some may have other intrinsic toxigenic material, such as cell wall teichoic acid, which can be immunostimulatory.

Many pathogenic bacteria excrete toxins into their environment (e.g., S. aureus enterotoxins); although there are no strict limits on concentrations of staphylococcal enterotoxins in medical products, such toxins could be present at clinically relevant concentrations in phage products. A strategy to avoid toxins produced by pathogenic bacteria would be to propagate phages in nonpathogenic species if possible (e.g., propagation of phages targeted at S. aureus using Staphylococcus xylosus [157], although this potentially reduces the number of available therapeutic phages, as not all phages possess such broad-range activity). Another approach is to assess phage preparation effects on viability of a eukaryotic cell line, although how this might be used to include or exclude a particular preparation is unclear (158).

Rationale.

As with antibiotics, there may be a need for accurate and reproducible methods that detect and quantify phages in complex clinical samples obtained from infection sites (84). Such data may be helpful to determine optimal dose, route of administration, dosing frequency, and treatment duration. However, few studies have assessed phage concentrations at infection sites. Ideally, assays that quantify phages in clinical specimens should be designed such that naturally present (nontherapeutic) phages are not detected.

The double agar overlay method and spot modification thereof (Question 7) may be reconfigured to quantitative assays to determine phage titers by using serial dilutions of known concentrations of phage as a comparator to unknown amounts of phage, with phage concentrations expressed as PFU per milliliter of the assayed preparation. This may underrepresent the number of active phages if a single plaque reflects activity of multiple phage particles, which, though theoretically possible, is likely rare. Likewise, liquid-based methods (Question 7) can be reconfigured to quantitative assays. Quantitative (including digital droplet) PCR can be used to enumerate phages based on detection of phage nucleic acid (84). In most cases, phage quantitative PCR platforms use probes for specificity (91, 159). Quantitative PCR may be sensitive and reproducible and, while accurate and potentially configurable as a high-throughput test, may be expensive and technically challenging. Upfront equipment costs, sample nucleic acid extraction, the need for specific primer and, if used, probe design for each phage species (or related phages) based on a priori sequencing results, and assay validation are among the complexities. Finally, quantitative PCR-based viral quantitation cannot discriminate between viable and nonviable phages and may therefore overestimate functional phage concentrations.

The ARLG Phage Taskforce was unable to recommend a standard method for phage enumeration in clinical specimens but favored the double agar overlay method as it measures viable phages, which is the primary interest with use of lytic phages. However, the members of the ARLG Phage Taskforce suggest that it is reasonable to perform both double-agar overlay and quantitative PCR on the same clinical specimen for enumeration of phage where possible, given that there is no clear gold standard. If a rapid high-throughput assay is needed to quantify a specific phage in multiple samples, the ARLG Phage Taskforce suggests that it may be possible to use quantitative PCR with the addition of a double-agar overlay method correction coefficient (if shown to be accurate) to translate results to viable phage counts. Even if correlations can be established, the ARLG Phage Taskforce suggests that it is prudent to continue to use the double-agar overlay method alongside quantitative PCR until the relationship between results of the two approaches and reproducibility of testing are established with certainty for the phage under investigation.

In addition to methods to quantify phage preliminarily introduced in Question 7 and above, phage particles have been enumerated in other ways. Transmission electron microscopy (TEM) after negative staining and epifluorescence microscopy after staining with DNA fluorochromes have been used to enumerate phage but are labor-intensive, time-consuming, and expensive. TEM is impractical for processing many samples concurrently and cannot be used for complex clinical samples (84, 160). Flow cytometry and nanoparticle detection by laser-illuminated optical microscopy, mass spectrometry, and next-generation sequencing also have been used but are likely unsuitable for routine use (84, 91, 161, 162).

New detection and enumeration methods should address diverse specimen types, ranging from blood, tissue, synovial fluid, and urine to cerebrospinal fluid; have a suitable turnaround time; be easy to use in clinical laboratories; address the myriad of phages likely to be used, and have an ability to detect more than one phage in the same sample (relevant to cocktail administration). Finally, the clinical utility of such assays would need to be demonstrated; further animal and clinical research studies are needed. To this end, there are no existing clinical data that demonstrate phage concentrations that predict optimal net bacterial killing at an infection site, with clinical outcome potentially dependent on many factors, including, but not limited to, administration route, bacterial burden, phage density, bacterial phage susceptibility, bacterial doubling time, presence of biofilm, efficiency of phage adsorption and infection, latency period (i.e., time required for phage replication), burst size (i.e., number of phages released from a single lysed bacterial cell), and phage removal rates (163,–166) (see Question 13).

If there is a critical need to determine phage concentrations in a specimen for clinical purposes, the ARLG Phage Taskforce suggests only measuring phage concentrations in patients with active infections, given the potential for self-amplification in the presence of bacteria. The ARLG Phage Taskforce was unable to identify the optimal time for phage enumeration postadministration but suggests a lag time between phage administration and quantification of at least a day to allow for self-amplification.

Rationale.

In clinical practice, lytic phages may be administered as a cocktail or a single phage (54). Cocktails may be preferred to maximize the number of phages that target a specific bacterium and/or to broaden the spectrum of bacterial activity (i.e., target multiple bacteria in polymicrobial infections). Cocktails are also a strategy to optimize bacterial killing over time and minimize the potential for resistance emergence, as component phages may target different bacterial receptors with independent mutations in each receptor required to achieve resistance (26, 163, 167, 168). However, the theoretical benefits of this approach to maximize net bacterial killing and minimize resistance development have not been substantiated and require clinical validation. A drawback with cocktails is that individual phages usually require a decrease in concentration when mixed into a single dose. There is also the potential that component phages agglomerate (169) or interfere with one another by competing for the same bacterial receptor or drive cross-resistance (51, 169,–172). Phages can be given sequentially, potentially avoiding antagonism among phages or driving of cross-resistance (51). Phages are often concurrently used with antibiotics, so antibiotic effects on phage therapy bear consideration.

Two primary paradigms exist to produce phage therapeutics: prebiomanufactured and on-demand biomanufactured based on individual patient need (173, 174). Prebiomanufactured phage products are prepared in advance and are available off the shelf for administration. Prebiomanufactured phage products are used for either empirical use (as discussed, empirical treatment is generally discouraged for compassionate use cases) or personalized use. Prebiomanufactured phage preparations can be personalized by selecting the most appropriate phages based on PST among an array of already-made phages (see limitations in Questions 6 and 7). Prebiomanufactured products may be comprised of one or many phages, with selection of phages based on the combined susceptibility of phages against a panel of representative clinical isolates, including multidrug-resistant isolates. In the United States, prebiomanufactured phage products are manufactured under current GMP regulations (175) and meet defined quality and uniformity specifications (see Question 10) (176).

It is possible to biomanufacture personalized phage products on demand; these are typically comprised of one or more phages with confirmed activity against an individual patient’s isolate(s) (54). With on-demand biomanufactured phage products, there is the potential to modify the phage product based on clinical response and/or emergence of phage resistance. The major drawback of on-demand biomanufactured phage products is the extended (i.e., weeks to months) associated manufacturing and regulatory timelines (177).

The Magistral Phage system is a regulatory framework developed in Belgium in which on-demand phages are prepared for individual patients in local compounding pharmacies (174). Typically, a pharmacist with a physician's prescription prepares such phage products by selecting phages from phage banks and incorporating them into a designated administration vehicle (174, 178). This decentralized strategy allows for nimble manufacturing of individualized therapies for patients as they need them (typically within hours to days), with no case-by-case approval required. There are some considerations with the use of magistral or compounded phage products. Phage selection criteria from a phage bank and appropriate library sizes have yet to be identified for compounded products (178). Preparations of magistral or compounded phage products do not have to necessarily adhere to GMP production requirements, as regulations for clinical production of phages have not been established in some Western countries (179).

Rationale.

Phages possess pharmacokinetic properties distinct from small-molecule antibiotics due to their large size, protein content, and self-replicating nature (180). Such size constraints may limit phage concentrations that can be attained per unit volume as well as phage uptake and transport across and into tissues in situ. Their protein-rich composition results in rapid elimination by the mononuclear phagocytic system and potential neutralization by antiphage antibodies (181, 182). Thus, phage concentrations at sites of infection, especially when systemically administered, are assumed to be substantially lower than the initial dose for most phage treatments before phage replication.

Preclinical and mathematical modeling studies suggest that factors responsible for net bacterial killing at the infection site include (i) density of bacterial cells (susceptible and resistant populations) and (ii) their growth rates, (iii) numbers of phages present, (iv) adsorption or infectivity rate of phages, (v) latency periods, and (vi) burst sizes and phage killing/removal/inactivation rates at sites of infection (163,–166). A special feature of phage therapy is the potential for low-dose phage treatments, as phages are biologically replicating therapeutic agents. In contrast to antibiotics, small numbers of phages may result in profound increases in phage densities at infection sites “if the target population of bacteria is sufficiently dense and physiologically, and genetically, amenable to phage replication” (163). Based on data generated from preclinical infection models, it has been proposed that target bacterial cell densities need to be over a replication or proliferation threshold of 10⁴ CFU/ml for self-amplification to occur (165, 183).

Numbers of phages at sites of infection achieved by self-amplification or passive treatment required for optimal net bacterial killing and phage resistance prevention have not been established. Currently, the actual multiplicity of infection (MOI_actual) ratio, which is the ratio of adsorbed phages to targeted bacteria, is the expression used to reflect the desired or targeted number of phages at infection sites relative to bacterial burden (164, 165). The target MOI_actual has been proposed to be 10 (165), with some clinical data suggesting that failure to achieve an MOI_actual of ≥10 is associated with treatment failure (11, 13, 16). Importantly, clinical trials data supporting the importance of this MOI_actual target are unavailable. Recent modeling data indicate that the MOI_actual should result in phage densities that far exceed the inundation threshold, which is the minimum phage concentration above which the bacterial population declines (184). When phage concentrations are substantially above the inundation threshold (i.e., inundative densities), phages are expected to infect all susceptible host cells in a relatively short time and bacterial counts to decline exponentially. Regardless of bacterial densities, several investigators suggest that a reasonable phage density capable of maximizing net bacterial killing at the infection site in a timely manner is 10⁸ PFU/ml (164, 166, 170, 184, 185).

It is important to note that reported MOI_actual and inundative phage density targets are based on mathematical modeling of preclinical infection model data (164,–166, 184). Mathematical and simulation models are useful in quantifying conditions that optimize net bacterial killing by phages at infection sites, but their utility is restricted by validity of model assumptions and input parameter values (186). For example, ratios of viable to nonviable phage particles and numbers of absorbed phages to numbers of bacteria at infection sites are difficult to quantify and likely variable over a treatment course, making it difficult to estimate the MOI_actual. Phages may bind bacterial debris from lysed bacteria and are heterogeneously distributed at infection sites, making it difficult to determine the actual number of active phages at an infection site that can infect bacterial cells (186, 187). Estimation of desired phage densities across modeling and simulation studies does not account for contributions of the immune system, bacterial host defense mechanisms [e.g., bacterial membrane vesicle production, which reduces phage virulence], concurrent receipt of antibiotics, and local host environments, all of which affect phage therapy outcomes (114, 185). Finally, modeling studies performed to support the development of phage therapy require clinical validation.

Rationale.

The route of administration is likely to affect outcomes of treatment since efficacy is dependent on phage concentrations at the site of infection. Common routes of administration for patients with acute and chronic infections include systemic injection (e.g., i.v., i.p., intramuscular, and subcutaneous), oral, and direct administration (e.g., topical, intra-articular, intravesicular, inhalation, and bladder irrigation) (see Question 1) (173, 181). Use of phage for prevention and treatment of biofilm formation on medical devices has also been studied (188). Selection of administration routes for patients with acute and chronic infections has been largely empirical, with most published clinical data describing i.v., direct administration, or oral administration of phages (173).

Systemic injection routes are efficient ways to deliver phages and treat intravascular as well as a broad range of infections (181). Following systemic injection, phages are typically observed in the circulation shortly after administration. Due to rapid clearance from the bloodstream by the mononuclear phagocyte system, systemic phage concentrations decline exponentially within 8 to 12h (189). Phages are also subject to inactivation by the complement system and circulating neutralizing antibodies (see Question 9) (181, 182). If phage therapy occurs over an extended period of time such that the generation of antiphage antibodies is possible, consideration should be given to adding or substituting a phage that does not cross-react serologically (185) (see Question 9). Generation of neutralizing antibodies appears to be most robust with the i.v. route of administration (182), which may be mitigated by specific dosing strategies (25). In addition to liver and spleen (part of the mononuclear phagocyte system), available data indicate that phages distribute into heart, skeletal muscles, bone marrow, thymus, kidneys, and bladder after systemic delivery and that their ability to concentrate in certain organs/systems is dose dependent. The lungs and brain are other organs where phages have been detected; “backwards” penetration of phages from the bloodstream to the gastrointestinal tract has also been demonstrated in animal studies (181).

Direct (e.g., intravesicular, intra-articular) or topical administration has been used to treat patients with PJI, VAD infection, osteomyelitis, and other infections (see Question 1). With systemic injections, only a fraction of the phage dose administered reaches nonvascular infection sites due to dilution, translocation inefficiencies, and phage loss or inactivation (190). In contrast, direct administration to infection sites may ensure a high titer of phages at the intended target site; however, results of direct administration have been mixed and best practices have not been established. One challenge with direct administration is that phages are nonmotile, proteinaceous particles that follow Bowman collision dynamics and have reduced efficacy when the selected route of administration does not facilitate timely and sustained distribution within spatially structured bacterial populations at the infection site (191). Another consideration is that components of direct administration applications may inactivate phages, limiting clinical utility (192).

Inhaled administration, a form of direct administration, has been used clinically for treatment of patients with respiratory tract infections, but the ability of inhaled phages to concentrate in anatomic components of the respiratory tract (e.g., bronchi, bronchioles, and alveoli) has not yet been systematically characterized in humans. In animal studies, phage titers achieved in the respiratory tract vary based on delivery method (e.g., inhaler, nebulization via jet, ultrasonic or vibrating mesh, aerosolization) (193,–195), device, formulation (e.g., liquid, freeze-drying powder, spray-drying powder, aerosol), and phage (194). When delivering phages via aerosolization, care must be taken to ensure that the aerosolization method chosen does not reduce phage viability (193,–195). Nevertheless, inhaled phage delivery, so long as active phages reach targeted bacteria at the infection site, may be an effective route of administration. Systems that aim to circumvent delivery challenges (e.g., controlled-release formulations) are under development (169, 171, 172).

Topical delivery has also been explored to deliver systemic titers of phages (196). Compared to systemic injection, direct or topical administration results in lower systemic concentrations, especially when applied to intact skin (181, 197). Further data with novel transdermal delivery systems are needed to determine future clinical utility.

Lastly, the oral route of administration has been long studied as a means for delivering phages both locally and systemically and is attractive due to convenience (181). Transcytosis, the transport of molecules from one side of a cell to the other via endocytosis and exocytosis, has been proposed as a mechanism for systemic absorption from gut epithelial cells (198). However, degrees of survival in the human gastrointestinal tract and systemic absorption are uncertain, and oral administration is a less efficient means for achieving systemic therapeutic phage concentrations relative to other administration routes. Factors that may affect passage through the gastrointestinal tract include acidity of the stomach, host immune responses, and inhibitory effects of intestinal mucins (95, 181). Recovery of phages in feces can be used as a marker of survival in the gastrointestinal tract. Strategies to enhance stability of phages at unfavorable pH or temperatures and avoid phage elimination from the body to maintain active phages at infective doses are being pursued (e.g., encapsulation of phages in various matrices) (199). To date, phages have been detected in blood, urine, and feces in a dose-dependent manner in animal models following oral phage administration (200, 201). However, recovery of phages was inconsistent and, in some cases, not detectable in blood (202).

Rationale.

Phage dosing, irrespective of route of administration, thus far has been largely empirical. In most published cases, phage formulations have been administered daily (8, 9, 21, 22, 24, 34, 45, 50), twice-daily (5, 8, 23, 25, 27, 35, 44, 47,–50, 52, 58, 62, 138), every 6 to 8 h (7, 20, 32, 36, 40, 46, 48, 54, 55, 57), or by extended infusion over hours (49) (Table 1). Iredell et al. detected phage DNA in blood up to 12h after dosing with staphylococcal phages in humans with bloodstream infections (56). This served, in part, as the rationale for 12-h dosing. Many patients are treated via more than one route of administration (e.g., i.v. and direct application) (5, 7, 44, 45, 49, 57, 58, 62, 138) (Table 1). Durations of therapy have spanned from 1 day (17, 37, 38, 41,–43, 50, 51, 64) to months (7, 8, 20, 23, 25, 34, 54, 62, 173) (Table 1). Limited studies in humans and animals show recovery of phages from feces and blood after oral administration to be dose and duration dependent (181). In vitro and animal direct administration studies have shown a spectrum of efficacy based on dose and duration (107, 203,–213). Independent of administration route, resistance to phages is more likely to occur with repeated dosing and prolonged treatment courses (14, 214, 215). While phage resistance emergence is undesirable, there are limited clinical data indicating that in some scenarios, bacteria may become more susceptible to antibiotics and less virulent when they develop phage resistance (9, 51, 54) (see Question 2).

Preparation of this document was funded in part by the National Institute of Allergy and Infectious Diseases of the National Institutes of Health under award number UM1 AI104681.

The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.